Manual of Clinical Microbiology, 11E (2015 - TRUEPDF).pdf

11 Manual of Clinical Microbiology TH EDITION

MCM11_FM.indd 1

3/19/15 1:07 PM

MCM11_FM.indd 2

3/19/15 1:07 PM

11 Manual of Clinical Microbiology TH EDITION

EDITORS IN CHIEF

JAMES H. JORGENSEN

MICHAEL A. PFALLER

Emeritus, Department of Pathology, University of Texas Health Science Center, San Antonio, Texas

T2 Biosystems, Lexington, Massachusetts, and Professor Emeritus, University of Iowa College of Medicine, Iowa City, Iowa

VOLUME EDITORS

KAREN C. CARROLL

MARIE LOUISE LANDRY

GUIDO FUNKE

SANDRA S. RICHTER

Department of Pathology, Division of Microbiology, The Johns Hopkins University School of Medicine, Baltimore, Maryland

Risch Laboratories Group, Schaan, Principality of Liechtenstein

Departments of Laboratory Medicine and Internal Medicine, Yale University School of Medicine, New Haven, Connecticut

Department of Laboratory Medicine, Cleveland Clinic, Cleveland, Ohio

DAVID W. WARNOCK

Faculty of Medical and Human Sciences, University of Manchester, Manchester, United Kingdom

Volume 1

MCM11_FM.indd 3

Washington, DC

3/19/15 1:07 PM

Copyright © 2015 by ASM Press. ASM Press is a registered trademark of the American Society for Microbiology. All rights reserved. No part of this publication may be reproduced or transmitted in whole or in part or reutilized in any form or by any means, electronic or mechanical, including photocopying and recording, or by any information storage and retrieval system, without permission in writing from the publisher. Disclaimer: To the best of the publisher’s knowledge, this publication provides information concerning the subject matter covered that is accurate as of the date of publication. The publisher is not providing legal, medical, or other professional services. Any reference herein to any specific commercial products, procedures, or services by trade name, trademark, manufacturer, or otherwise does not constitute or imply endorsement, recommendation, or favored status by the American Society for Microbiology (ASM). The views and opinions of the author(s) expressed in this publication do not necessarily state or reflect those of ASM, and they shall not be used to advertise or endorse any product.

Library of Congress Cataloging-in-Publication Data Manual of clinical microbiology / editors in chief, James H. Jorgensen, Department of Pathology, University of Texas Health Science Center, San Antonio, Texas, Michael A. Pfaller, T2 Biosystems, Lexington, Massachusetts, and Professor Emeritus, University of Iowa College of Medicine, Iowa City, Iowa ; volume editors, Karen C. Carroll, Department of Pathology, Division of Microbiology, The Johns Hopkins Hospital, Baltimore, Maryland [and 4 others]. — 11th edition. volumes cm Includes bibliographical references and index. ISBN 978-1-55581-737-4 print set — ISBN 978-1-55581-738-1 e-book set 1. Medical microbiology—Handbooks, manuals, etc. 2. Diagnostic microbiology—Handbooks, manuals, etc. I. Jorgensen, James H., editor. II. Pfaller, Michael A., editor. III. Carroll, Karen C., editor. IV. American Society for Microbiology, issuing body. QR46.M425 2015 616.9'041—dc23 2015008089

doi:10.1128/9781555817381 Printed in Canada 10 9 8 7 6 5 4 3 2 1 Address editorial correspondence to: ASM Press, 1752 N St., N.W., Washington, DC 20036-2904, USA. Send orders to: ASM Press, P.O. Box 605, Herndon, VA 20172, USA. Phone: 800-546-2416; 703-661-1593. Fax: 703-661-1501. E-mail: [email protected] Online: http://estore.asm.org

MCM11_FM.indd 4

4/2/15 1:09 AM

Contents

Volume 1

8 Prevention of Health Care-Associated Infections / 106

DANIEL J. DIEKEMA AND MICHAEL A. PFALLER

9 Investigation of Disease Outbreaks / 120

Editorial Board xi

IONA MUNJAL AND BELINDA OSTROWSKY

Contributors xiii

10 Molecular Epidemiology / 131

EIJA TREES, PAUL A. ROTA, DUNCAN MacCANNELL, AND PETER GERNER-SMIDT

Preface xxv Author and Editor Conflicts of Interest xxvi

11 Procedures for the Storage of Microorganisms / 161

I

ROSEMARY C. SHE AND CATHY A. PETTI

s ec tion Diagnostic strategies and general topics / 1

12 Prevention of Laboratory-Acquired Infections / 169 MICHAEL A. NOBLE

13 Decontamination, Disinfection, and Sterilization / 183

VOLUME EDITOR: SANDRA S. RICHTER

CONSTANZE WENDT, RENO FREI, AND ANDREAS F. WIDMER

SECTION EDITOR: ROBIN PATEL

1

14 Biothreat Agents / 217

Introduction to the 11th Edition of the Manual of Clinical Microbiology / 3

SUSAN E. SHARP AND MICHAEL LOEFFELHOLZ

15 The Human Microbiome / 226

JAMES H. JORGENSEN AND MICHAEL A. PFALLER

2

Microscopy / 5

3

Laboratory Detection of Bacteremia and Fungemia / 15

JAMES VERSALOVIC, SARAH K. HIGHLANDER, AND JOSEPH F. PETROSINO

16 Microbial Genomics and Pathogen Discovery / 238

DANNY L. WIEDBRAUK

JENNIFER K. SPINLER, PEERA HEMARAJATA, AND JAMES VERSALOVIC

MICHAEL L. WILSON, MELVIN P. WEINSTEIN, AND L. BARTH RELLER

4

II

Systems for Identification of Bacteria and Fungi / 29

s ec tion Bacteriology / 252

KAREN C. CARROLL AND ROBIN PATEL

5

Automation and Design of the Clinical Microbiology Laboratory / 44 CHRISTOPHER D. DOERN AND MARTIN HOLFELDER

6

Molecular Microbiology / 54

7

Immunoassays for Diagnosis of Infectious Diseases / 91

VOLUME EDITORS: KAREN C. CARROLL AND GUIDO FUNKE SECTION EDITORS: KATHRYN A. BERNARD, J. STEPHEN DUMLER, MELISSA B. MILLER, CATHY A. PETTI, AND PETER A. R. VANDAMME

FREDERICK S. NOLTE

General

ELITZA S. THEEL, A. BETTS CARPENTER, AND MATTHEW J. BINNICKER

17 Taxonomy and Classification of Bacteria / 255 PETER A. R. VANDAMME

v

MCM11_FM.indd 5

3/19/15 1:07 PM

vi

n  CONTENTS

18 Specimen Collection, Transport, and Processing: Bacteriology / 270 ELLEN JO BARON

19 Reagents, Stains, and Media: Bacteriology / 316 RONALD ATLAS AND JAMES SNYDER

Gram-Positive Cocci

32 Mycobacterium: Clinical and Laboratory Characteristics of Rapidly Growing Mycobacteria / 595 BARBARA A. BROWN-ELLIOTT AND RICHARD J. WALLACE, JR.

Gram-Negative Bacteria

20 General Approaches to Identification of Aerobic Gram-Positive Cocci / 350

33 Approaches to the Identification of Aerobic Gram-Negative Bacteria / 613

21 Staphylococcus, Micrococcus, and Other Catalase-Positive Cocci / 354

34 Neisseria / 635

JENS JØRGEN CHRISTENSEN AND KATHRYN L. RUOFF

KARSTEN BECKER, ROBERT L. SKOV, AND CHRISTOF von EIFF

GEORGES WAUTERS AND MARIO VANEECHOUTTE

JOHANNES ELIAS, MATTHIAS FROSCH, AND ULRICH VOGEL

22 Streptococcus / 383

35 Aggregatibacter, Capnocytophaga, Eikenella, Kingella, Pasteurella, and Other Fastidious or Rarely Encountered Gram-Negative Rods / 652

23 Enterococcus / 403

36 Haemophilus / 667

BARBARA SPELLERBERG AND CLAUDIA BRANDT LÚCIA MARTINS TEIXEIRA, MARIA DA GLÓRIA SIQUEIRA CARVALHO, RICHARD R. FACKLAM, AND PATRICIA LYNN SHEWMAKER

24 Aerococcus, Abiotrophia, and Other Aerobic Catalase-Negative, Gram-Positive Cocci / 422 JENS JØRGEN CHRISTENSEN AND KATHRYN L. RUOFF

Gram-Positive Rods

REINHARD ZBINDEN

NATHAN A. LEDEBOER AND GARY V. DOERN

37 Escherichia, Shigella, and Salmonella / 685 NANCY A. STROCKBINE, CHERYL A. BOPP, PATRICIA I. FIELDS, JAMES B. KAPER, AND JAMES P. NATARO

38 Klebsiella, Enterobacter, Citrobacter, Cronobacter, Serratia, Plesiomonas, and Other Enterobacteriaceae / 714 STEPHEN J. FORSYTHE, SHARON L. ABBOTT, AND JOHANN PITOUT

25 General Approaches to the Identification of Aerobic Gram-Positive Rods / 437

39 Yersinia / 738

26 Bacillus and Other Aerobic Endospore-Forming Bacteria / 441

40 Aeromonas / 752

KATHRYN A. BERNARD

CHRISTINE Y. TURENNE, JAMES W. SNYDER, AND DAVID C. ALEXANDER

27 Listeria and Erysipelothrix / 462 NELE WELLINGHAUSEN

28 Coryneform Gram-Positive Rods / 474

GUIDO FUNKE AND KATHRYN A. BERNARD

29 Nocardia, Rhodococcus, Gordonia, Actinomadura, Streptomyces, and Other Aerobic Actinomycetes / 504 PATRICIA S. CONVILLE AND FRANK G. WITEBSKY

30 Mycobacterium: General Characteristics, Laboratory Detection, and Staining Procedures / 536 GABY E. PFYFFER

31 Mycobacterium: Laboratory Characteristics of Slowly Growing Mycobacteria / 570

PATRICIA J. SIMNER, STEFFEN STENGER, ELVIRA RICHTER, BARBARA A. BROWN-ELLIOTT, RICHARD J. WALLACE, JR., AND NANCY L.WENGENACK

JEANNINE M. PETERSEN, LORI M. GLADNEY, AND MARTIN E. SCHRIEFER AMY J. HORNEMAN

41 Vibrio and Related Organisms / 762

CHERYL L. TARR, CHERYL A. BOPP, AND J. J. FARMER, III

42 Pseudomonas / 773

NIELS HØIBY, OANA CIOFU, AND THOMAS BJARNSHOLT

43 Burkholderia, Stenotrophomonas, Ralstonia, Cupriavidus, Pandoraea, Brevundimonas, Comamonas, Delftia, and Acidovorax / 791 JOHN J. LiPUMA, BART J. CURRIE, SHARON J. PEACOCK, AND PETER A. R. VANDAMME

44 Acinetobacter, Chryseobacterium, Moraxella, and Other Nonfermentative Gram-Negative Rods / 813 MARIO VANEECHOUTTE, ALEXANDR NEMEC, PETER KÄMPFER, PIET COOLS, AND GEORGES WAUTERS

45 Bordetella and Related Genera / 838

CARL-HEINZ WIRSING von KÖNIG, MARION RIFFELMANN, AND TOM COENYE

46 Francisella / 851

JEANNINE M. PETERSEN AND MARTIN E. SCHRIEFER

MCM11_FM.indd 6

3/19/15 1:07 PM

CONTENTS

n  vii

47 Brucella / 863

62 Mycoplasma and Ureaplasma / 1088

48 Bartonella / 873

63 Chlamydiaceae / 1106

49 Legionella / 887

64 Rickettsia and Orientia / 1122

Anaerobic Bacteria

65 Ehrlichia, Anaplasma, and Related Intracellular Bacteria / 1135

50 Approaches to Identification of Anaerobic Bacteria / 905

66 Coxiella / 1150

GEORGE F. ARAJ DIANA G. SCORPIO AND J. STEPHEN DUMLER PAUL H. EDELSTEIN AND CHRISTIAN LÜCK

ELLEN JO BARON

51 Peptostreptococcus, Finegoldia, Anaerococcus, Peptoniphilus, Veillonella, and Other Anaerobic Cocci / 909

KEN B. WAITES AND DAVID TAYLOR-ROBINSON CHARLOTTE A. GAYDOS AND ANDREAS ESSIG DAVID H. WALKER AND DONALD H. BOUYER

MEGAN E. RELLER AND J. STEPHEN DUMLER

STEPHEN R. GRAVES AND ROBERT F. MASSUNG

67 Tropheryma whipplei / 1159

WALTER GEIßDÖRFER, ANNETTE MOTER, AND CHRISTIAN BOGDAN

YULI SONG AND SYDNEY M. FINEGOLD

52 Propionibacterium, Lactobacillus, Actinomyces, and Other Non-Spore-Forming Anaerobic Gram-Positive Rods / 920 VAL HALL AND SARAH D. COPSEY

53 Clostridium / 940

DENNIS L. STEVENS, AMY E. BRYANT, AND KAREN C. CARROLL

54 Bacteroides, Porphyromonas, Prevotella, Fusobacterium, and Other Anaerobic Gram-Negative Rods / 967 EIJA KÖNÖNEN, GEORG CONRADS, AND ELISABETH NAGY

Curved and Spiral-Shaped Gram-Negative Rods 55 Algorithms for Identification of Curved and Spiral-Shaped Gram-Negative Rods / 994 IRVING NACHAMKIN

56 Campylobacter and Arcobacter / 998 COLLETTE FITZGERALD AND IRVING NACHAMKIN

57 Helicobacter / 1013 ANDY J. LAWSON

58 Leptospira / 1028 PAUL N. LEVETT

59 Borrelia / 1037

MARTIN E. SCHRIEFER

60 Treponema and Brachyspira, Human Host-Associated Spirochetes / 1055

ARLENE C. SEÑA, ALLAN PILLAY, DAVID L. COX, AND JUSTIN D. RADOLF

III

s ec tion Antibacterial Agents and Susceptibility Test Methods / 1169 VOLUME EDITOR: SANDRA S. RICHTER SECTION EDITOR: JEAN B. PATEL

68 Antibacterial Agents / 1171

JAMES S. LEWIS, II, AND KAREN BUSH

69 Mechanisms of Resistance to Antibacterial Agents / 1212 JEAN B. PATEL AND SANDRA S. RICHTER

70 Susceptibility Test Methods: General Considerations / 1246 JOHN D. TURNIDGE

71 Susceptibility Test Methods: Dilution and Disk Diffusion Methods / 1253

JAMES H. JORGENSEN AND JOHN D. TURNIDGE

72 Antimicrobial Susceptibility Testing Systems / 1274 JAMES A. KARLOWSKY AND SANDRA S. RICHTER

73 Special Phenotypic Methods for Detecting Antibacterial Resistance / 1286

BRANDI M. LIMBAGO AND JANA M. SWENSON

74 Susceptibility Test Methods: Fastidious Bacteria / 1314 ROMNEY M. HUMPHRIES AND JANET A. HINDLER

Mycoplasmas and Obligate Intracellular Bacteria

75 Susceptibility Test Methods: Anaerobic Bacteria / 1342

61 General Approaches to Identification of Mycoplasma, Ureaplasma, and Obligate Intracellular Bacteria / 1082

76 Susceptibility Test Methods: Mycobacteria, Nocardia, and Other Actinomycetes / 1356

J. STEPHEN DUMLER

MCM11_FM.indd 7

AUDREY N. SCHUETZ AND DAVID W. HECHT

GAIL L. WOODS, SHOU-YEAN GRACE LIN, AND EDWARD P. DESMOND

3/19/15 1:07 PM

viii

n  CONTENTS

77 Molecular Detection of Antibacterial Drug Resistance / 1379 APRIL N. ABBOTT AND FERRIC C. FANG

Author Index xxix Subject Index xxxi

86 Respiratory Syncytial Virus and Human Metapneumovirus / 1498 N. ESTHER BABADY AND YI-WEI TANG

87 Measles and Rubella Viruses / 1519

WILLIAM J. BELLINI AND JOSEPH P. ICENOGLE

88 Enteroviruses and Parechoviruses / 1536 KATHLEEN A. STELLRECHT, DARYL M. LAMSON, AND JOSÉ R. ROMERO

Volume 2

89 Rhinoviruses / 1551

Editorial Board xi

91 Hepatitis A and E Viruses / 1584

Contributors xiii Preface xxv Author and Editor Conflicts of Interest xxvi

IV

s ec tion Virology / 1390

MARIE LOUISE LANDRY AND XIAOYAN LU

90 Coronaviruses / 1565

NAOMI J. GADSBY AND KATE E. TEMPLETON DAVID A. ANDERSON AND NATALIE A. COUNIHAN

92 Hepatitis C Virus / 1599

MICHAEL S. FORMAN AND ALEXANDRA VALSAMAKIS

93 Gastroenteritis Viruses / 1617

XIAOLI PANG AND RICHARD L. HODINKA

94 Rabies Virus / 1633

LILLIAN A. ORCIARI, CATHLEEN A. HANLON, AND RICHARD FRANKA

VOLUME EDITOR: MARIE LOUISE LANDRY

95 Arboviruses / 1644

SECTION EDITORS: ANGELA M. CALIENDO, CHRISTINE C. GINOCCHIO, YI-WEI TANG, AND ALEXANDRA VALSAMAKIS

96 Hantaviruses / 1660

General 78 Taxonomy and Classification of Viruses / 1393 ELLIOT J. LEFKOWITZ

79 Specimen Collection, Transport, and Processing: Virology / 1405 JAMES J. DUNN

80 Reagents, Stains, Media, and Cell Cultures: Virology / 1422

CHRISTINE C. GINOCCHIO, GERALD VAN HORN, AND PATRICIA HARRIS

81 Algorithms for Detection and Identification of Viruses / 1432

MARIE LOUISE LANDRY, ANGELA M. CALIENDO, CHRISTINE C. GINOCCHIO, YI-WEI TANG, AND ALEXANDRA VALSAMAKIS

ELIZABETH HUNSPERGER CHARLES F. FULHORST AND MICHAEL D. BOWEN

97 Arenaviruses and Filoviruses / 1669

PIERRE E. ROLLIN, STUART T. NICHOL, SHERIF ZAKI, AND THOMAS G. KSIAZEK

DNA Viruses 98 Herpes Simplex Viruses and Herpes B Virus / 1687 KEITH R. JEROME AND RHODA ASHLEY MORROW

99 Varicella-Zoster Virus / 1704

ELISABETH PUCHHAMMER-STÖCKL AND STEPHAN W. ABERLE

100 Human Cytomegalovirus / 1718 RICHARD L. HODINKA

101 Epstein-Barr Virus / 1738

RNA Viruses 82 Human Immunodeficiency Viruses / 1436

BERNARD M. BRANSON AND S. MICHELE OWEN

83 Human T-Cell Lymphotropic Viruses / 1458

WILLIAM M. SWITZER, WALID HENEINE, AND S. MICHELE OWEN

84 Influenza Viruses / 1470

ROBERT L. ATMAR AND STEPHEN E. LINDSTROM

85 Parainfluenza and Mumps Viruses / 1487 DIANE S. LELAND

MCM11_FM.indd 8

BARBARA C. GÄRTNER AND JUTTA PREIKSAITIS

102 Human Herpesviruses 6, 7, and 8 / 1754 PHILIP E. PELLETT AND GRAHAM TIPPLES

103 Adenoviruses / 1769

MARCELA ECHAVARRIA, CHRISTINE ROBINSON, AND RANDALL T. HAYDEN

104 Human Papillomaviruses / 1783

CHRISTINE C. GINOCCHIO, PATTI E. GRAVITT, AND JENNIFER S. SMITH

3/19/15 1:07 PM

CONTENTS n  ix

105 Human Polyomaviruses / 1803 RICHARD S. BULLER

106 Parvovirus B19 and Bocaviruses / 1818

115 Reagents, Stains, and Media: Mycology / 1955 MARK D. LINDSLEY, JAMES W. SNYDER, RONALD M. ATLAS, AND MARK T. LaROCCO

107 Poxviruses / 1828

116 General Approaches for Direct Detection and Identification of Fungi / 1965

108 Hepatitis B and D Viruses / 1841

Fungi

KEVIN E. BROWN

LAURA HUGHES, VICTORIA A. OLSON, AND INGER K. DAMON REBECCA T. HORVAT AND RYAN TAYLOR

Subviral Agents 109 Transmissible Spongiform Encephalopathies / 1859

MARKUS GLATZEL AND ADRIANO AGUZZI

V

s ec tion Antiviral Agents and Susceptibility Test Methods / 1867 VOLUME EDITOR: MARIE LOUISE LANDRY SECTION EDITORS: ANGELA M. CALIENDO, CHRISTINE C. GINOCCHIO, AND ALEXANDRA VALSAMAKIS

110 Antiviral Agents / 1869

AIMEE C. HODOWANEC, KENNETH D. THOMPSON, AND NELL S. LURAIN

111 Mechanisms of Resistance to Antiviral Agents / 1894

ROBERT W. SHAFER AND SUNWEN CHOU

112 Susceptibility Test Methods: Viruses / 1913 DIANA D. HUANG AND MATTHEW J. BANKOWSKI

VI

H. RUTH ASHBEE

117 Candida, Cryptococcus, and Other Yeasts of Medical Importance / 1984 SUSAN A. HOWELL, KEVIN C. HAZEN, AND MARY E. BRANDT

118 Pneumocystis / 2015 MELANIE T. CUSHION

119 Aspergillus and Penicillium / 2030

SHARON C.-A. CHEN, TANIA C. SORRELL, AND WIELAND MEYER

120 Fusarium and Other Opportunistic Hyaline Fungi / 2057 SEAN X. ZHANG, KERRY O’DONNELL, AND DEANNA A. SUTTON

121 Agents of Systemic and Subcutaneous Mucormycosis and Entomophthoromycosis / 2087

DEA GARCIA-HERMOSO, ALEXANDRE ALANIO, OLIVIER LORTHOLARY, AND FRANÇOISE DROMER

122 Histoplasma, Blastomyces, Coccidioides, and Other Dimorphic Fungi Causing Systemic Mycoses / 2109 GEORGE R. THOMPSON III AND BEATRIZ L. GÓMEZ

123 Trichophyton, Microsporum, Epidermophyton, and Agents of Superficial Mycoses / 2128 ANDREW M. BORMAN AND RICHARD C. SUMMERBELL

124 Curvularia, Exophiala, Scedosporium, Sporothrix, and Other Melanized Fungi / 2153 JOSEP GUARRO AND G. SYBREN de HOOG

s ec tion Mycology / 1932

125 Fungi Causing Eumycotic Mycetoma / 2173

VOLUME EDITOR: DAVID W. WARNOCK

126 Mycotoxins / 2188

SECTION EDITORS: MARY E. BRANDT AND ELIZABETH M. JOHNSON

General 113 Taxonomy and Classification of Fungi / 1935 MARY E. BRANDT AND DAVID W. WARNOCK

114 Specimen Collection, Transport, and Processing: Mycology / 1944 KARIN L. McGOWAN

MCM11_FM.indd 9

ABDALLA O. A. AHMED, G. SYBREN de HOOG, AND WENDY W. J. van de SANDE KURT THROCKMORTON, NANCY C. ISHAM, MAHMOUD A. GHANNOUM, AND NANCY KELLER

127 Lacazia, Lagenidium, Pythium, and Rhinosporidium / 2196

RAQUEL VILELA AND LEONEL MENDOZA

128 Microsporidia / 2209

RAINER WEBER, PETER DEPLAZES, AND ALEXANDER MATHIS

3/19/15 1:07 PM

x

n  CONTENTS

VII

s ec tion Antifungal Agents and Susceptibility Test Methods / 2221

139 Pathogenic and Opportunistic Free-Living Amebae / 2387 GOVINDA S. VISVESVARA

140 Intestinal and Urogenital Amebae, Flagellates, and Ciliates / 2399 SUSAN NOVAK-WEEKLEY AND AMY L. LEBER

VOLUME EDITOR: DAVID W. WARNOCK

141 Cystoisospora, Cyclospora, and Sarcocystis / 2425

SECTION EDITORS: MARY E. BRANDT AND ELIZABETH M. JOHNSON

142 Cryptosporidium / 2435

129 Antifungal Agents / 2223

SHAWN R. LOCKHART AND DAVID W. WARNOCK

130 Mechanisms of Resistance to Antifungal Agents / 2236 DAVID S. PERLIN

131 Susceptibility Test Methods: Yeasts and Filamentous Fungi / 2255 ELIZABETH M. JOHNSON AND MAIKEN CAVLING-ARENDRUP

VIII

s ec tion Parasitology / 2282 VOLUME EDITOR: DAVID W. WARNOCK SECTION EDITORS: BOBBI S. PRITT AND GARY W. PROCOP

General 132 Taxonomy and Classification of Human Parasitic Protozoa and Helminths / 2285 FRANCIS E. G. COX

133 Specimen Collection, Transport, and Processing: Parasitology / 2293

ROBYN Y. SHIMIZU AND LYNNE S. GARCIA

134 Reagents, Stains, and Media: Parasitology / 2310

ANDREA J. LINSCOTT AND SUSAN E. SHARP

135 General Approaches for Detection and Identification of Parasites / 2317

LYNNE S. GARCIA, GRAEME P. PALTRIDGE, AND ROBIN Y. SHIMIZU

Parasites 136 Plasmodium and Babesia / 2338 BOBBI S. PRITT

DAVID S. LINDSAY AND LOUIS M. WEISS LIHUA XIAO AND VITALIANO CAMA

143 Nematodes / 2448

HARSHA SHEOREY, BEVERLEY-ANN BIGGS, AND NORBERT RYAN

144 Filarial Nematodes / 2461

SOUMYA CHATTERJEE AND THOMAS B. NUTMAN

145 Cestodes / 2471

HECTOR H. GARCIA, JUAN A. JIMENEZ, AND HERMES ESCALANTE

146 Trematodes / 2479

MALCOLM K. JONES, JENNIFER KEISER, AND DONALD P. MCMANUS

147 Less Common Helminths / 2493

GARY W. PROCOP AND RONALD C. NEAFIE

148 Arthropods of Medical Importance / 2505 SAM R. TELFORD III

IX

s ec tion Antiparasitic Agents and Susceptibility Test Methods / 2527 VOLUME EDITOR: DAVID W. WARNOCK SECTION EDITOR: GARY W. PROCOP

149 Antiparasitic Agents / 2529

KARIN LEDER AND PETER F. WELLER

150 Mechanisms of Resistance to Antiparasitic Agents / 2550 W. EVAN SECOR, JACQUES LE BRAS, AND JÉRÔME CLAIN

151 Susceptibility Test Methods: Parasites / 2563 JACQUES LE BRAS, JÉRÔME CLAIN, AND W. EVAN SECOR

137 Leishmania and Trypanosoma / 2357

Author Index xxix

138 Toxoplasma / 2373

Subject Index xxxi

DAVID A. BRUCKNER AND JAIME LABARCA JAMES B. McAULEY, JEFFREY L. JONES, AND KAMALJIT SINGH

MCM11_FM.indd 10

3/19/15 1:07 PM

Editorial Board

Kathryn A. Bernard

Section II

Robin Patel

National Microbiology Laboratory, Public Health Agency of Canada, Winnipeg, Manitoba R3E 3R2, Canada

Mary E. Brandt

Sections VI and VII

Sections IV and V

Yi-Wei Tang

Sections IV and V

Sections VI and VII

Mycology Reference Laboratory, HPA South West Laboratory, Kingsdown, Bristol BS2 8EL, United Kingdom

Section IV

Alexandra Valsamakis

Sections IV and V

Peter A. R. Vandamme

Section II

Division of Medical Microbiology, Department of Pathology, The Johns Hopkins Hospital, Baltimore, MD 21287-7093

Section II

Department of Pathology and Laboratory Medicine, University of North Carolina School of Medicine, Chapel Hill, NC 27599-7525

Jean B. Patel

Sections VIII and IX

Department of Laboratory Medicine, Memorial SloanKettering Cancer Center, and Department of Pathology and Laboratory Medicine, Weill Medical College of Cornell University, New York, NY 10065

Hofstra North Shore-LIJ School of Medicine, Hempstead, NY 11549, and bioMérieux, Durham, NC 27712

Melissa B. Miller

Section VIII

Department of Molecular Pathology, Cleveland Clinic, Pathology and Laboratory Medicine Institute, Cleveland, OH 44195

Departments of Pathology and Microbiology & Immunology, University of Maryland School of Medicine, Baltimore, MD 21201

Elizabeth M. Johnson

Bobbi S. Pritt

Gary W. Procop

Section II

Christine C. Ginocchio

Section II

Department of Laboratory Medicine and Pathology, Division of Clinical Microbiology, Mayo Clinic, Rochester, MN 55905

Department of Medicine, Warren Alpert Medical School of Brown University, Providence, RI 02903

J. Stephen Dumler

Cathy A. Petti

HealthSpring Global Inc., Bradenton, FL 34209

Mycotic Diseases Branch, Centers for Disease Control and Prevention, Atlanta, GA 30333

Angela M. Caliendo

Section I

Division of Clinical Microbiology, Mayo Clinic, Rochester, MN 55905

Department of Biochemistry and Microbiology, Laboratory of Microbiology, Universiteit Gent, B-9000 Gent, Belgium

Section III

Office of Antimicrobial Resistance, Division of Healthcare Quality Promotion, Centers for Disease Control and Prevention, Atlanta, GA 30333 xi

MCM11_FM.indd 11

3/19/15 1:07 PM

MCM11_FM.indd 12

3/19/15 1:07 PM

Contributors

APRIL N. ABBOTT

H. RUTH ASHBEE

Department of Laboratory Medicine, University of Washington School of Medicine, Seattle, WA 98195

Mycology Reference Centre, Department of Microbiology, Leeds Teaching Hospitals NHS Trust, Leeds, LS1 3EX, United Kingdom

SHARON L. ABBOTT

Microbial Diseases Laboratory, Division of Communicable Disease Control, California Department of Public Health, Richmond, CA 94804

RONALD M. ATLAS

STEPHAN W. ABERLE

ROBERT L. ATMAR

Department of Biology, University of Louisville, Louisville, KY 40292

Department of Virology, Medical University of Vienna, Kinderspitalgasse 15, A-1095 Vienna, Austria

Departments of Medicine and Molecular Virology & Microbiology, Baylor College of Medicine, 1 Baylor Plaza, MS BCM280, Houston, TX 77030

ADRIANO AGUZZI

N. ESTHER BABADY

Institute of Neuropathology, University Hospital of Zürich, Schmelzbergstrasse 12, CH-8091 Zürich, Switzerland

Department of Laboratory Medicine, Memorial SloanKettering Cancer Center, New York, NY 10065

ABDALLA O. A. AHMED

MATTHEW J. BANKOWSKI

Department of Medical Microbiology, Faculty of Medicine, Umm Al-Qura University, Prince Sultan Street, 21955 Makkah Al Mukarramah, Saudi Arabia

Diagnostic Laboratory Services, Inc. (Queen’s Medical Center), Aiea, Hawaii, and Dept. of Pathology and Dept. of Tropical Medicine, Medical Microbiology and Pharmacology, John A. Burns School of Medicine, University of Hawaii at Manoa, Honolulu, HI 96813

ALEXANDRE ALANIO

Unité Mycologie Moléculaire, CNRS URA3012, Centre National de Référence des Mycoses Invasives et Antifongiques, Institut Pasteur, Paris, France

ELLEN JO BARON

Emerita, Stanford University, and Department of Technical Support, Cepheid, 904 Caribbean Drive, Sunnyvale, CA 94089

DAVID C. ALEXANDER

Saskatchewan Disease Control Laboratory, Saskatchewan Ministry of Health, Regina, Saskatchewan S4S 0A4, Canada

KARSTEN BECKER

University Hospital Münster, Institute of Medical Microbiology, D-48149 Münster, Germany

DAVID A. ANDERSON

Macfarlane Burnet Institute for Medical Research and Public Health, AMREP, 85 Commercial Road, Melbourne 3004, Australia

WILLIAM J. BELLINI

Measles, Mumps, Rubella and Herpesviruses Laboratory Branch, Mail Stop C-22, Centers for Disease Control and Prevention, 1600 Clifton Road, NE, Atlanta, GA 30333

GEORGE F. ARAJ

Department of Pathology & Laboratory Medicine, American University of Beirut Medical Center, PO Box 11-0236, Beirut, Lebanon 1107-2020

KATHRYN A. BERNARD

National Microbiology Laboratory, Public Health Agency of Canada, 1015 Arlington St., Winnipeg, MB R3E 3R2, Canada

xiii

MCM11_FM.indd 13

3/19/15 1:07 PM

xiv

n  CONTRIBUTORS

BEVERLEY-ANN BIGGS

DAVID A. BRUCKNER

MATTHEW J. BINNICKER

AMY E. BRYANT

THOMAS BJARNSHOLT

RICHARD S. BULLER

Victorian Infectious Diseases Service, Royal Melbourne Hospital, and Department of Medicine, University of Melbourne, Parkville, Victoria 3050, Australia Department of Laboratory Medicine and Pathology, Division of Clinical Microbiology, Mayo Clinic, Rochester, MN 55901 Department of Clinical Microbiology, Rigshospitalet, and Department of Immunology and Microbiology, Faculty of Health Sciences, University of Copenhagen, Copenhagen, Denmark

CHRISTIAN BOGDAN

Mikrobiologisches Institut - Klinische Mikrobiologie, Immunologie und Hygiene, Friedrich-Alexander-Universität Erlangen-Nürnberg and Universitätsklinikum Erlangen, 91054 Erlangen, Germany

CHERYL A. BOPP

Enteric Diseases Laboratory Branch, Centers for Disease Control and Prevention, Atlanta, GA 30333

ANDREW M. BORMAN

United Kingdom National Mycology Reference Laboratory, Public Health England, Myrtle Road, Bristol, BS2 8EL, United Kingdom

DONALD H. BOUYER

Department of Pathology, University of Texas Medical Branch, Galveston, TX 77555-0609

MICHAEL D. BOWEN

Gastroenteritis and Respiratory Viruses Laboratory Branch, Division of Viral Diseases, NCIRD, Centers for Disease Control and Prevention, Atlanta, GA 30333

CLAUDIA BRANDT

Institute of Medical Microbiology, Johann Wolfgang Goethe University, Paul Ehrlich Str. 40, 60596 Frankfurt, Germany

MARY E. BRANDT

Mycotic Diseases Branch, Centers for Disease Control and Prevention, 1600 Clifton Road, Mailstop G-11, Atlanta, GA 30333

BERNARD M. BRANSON

Division of HIV/AIDS Prevention, National Center for HIV/ AIDS, Viral Hepatitis, STD, and TB Prevention, Centers for Disease Control and Prevention, Atlanta, GA 30333

KEVIN E. BROWN

Virus Reference Department, Public Health England, London, NW9 5EQ, United Kingdom

BARBARA A. BROWN-ELLIOTT

University of Texas Health Science Center at Tyler, Department of Microbiology, 11937 U.S. Highway 271, Tyler, TX 75708

MCM11_FM.indd 14

Dept. of Pathology & Laboratory Medicine, David Geffen UCLA School of Medicine, 3741 Mountain View Ave., Los Angeles, CA 90066-3111 Veterans Affairs Medical Center, Boise, ID 83702, and University of Washington School of Medicine, Seattle, WA 98195 Department of Pediatrics, Box 8116, Washington University School of Medicine, 660 S. Euclid Ave., St. Louis, MO 63110

KAREN BUSH

Departments of Molecular and Cellular Biochemistry and Biology, Indiana University, Bloomington, IN 47405

ANGELA M. CALIENDO

Department of Medicine, Warren Alpert Medical School of Brown University, Providence, RI 02903

VITALIANO CAMA

CGH/Div. of Parasitic Diseases and Malaria/Parasitic Diseases Branch, Centers for Disease Control and Prevention, 1600 Clifton Road NE, Atlanta, GA 30329-4018

A. BETTS CARPENTER

Molecular Microbiology & Cytology Laboratories, LabCorp, and Department of Pathology, Joan C. Edwards School of Medicine, Marshall University, Huntington, WV 25704

KAREN C. CARROLL

Department of Pathology, Division of Microbiology, The Johns Hopkins University School of Medicine, Baltimore, MD 21287

MARIA DA GLÓRIA SIQUEIRA CARVALHO

Streptococcus Laboratory, Respiratory Diseases Branch, Division of Bacterial Diseases, Centers for Disease Control and Prevention, Mail Stop C0-2, Atlanta, GA 30333

MAIKEN CAVLING-ARENDRUP

Dept. of Microbiology & Infection Control, Statens Serum Institut, Artillerivej 5, DK-2300 Copenhagen, Denmark

SOUMYA CHATTERJEE

Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892

SHARON C.-A. CHEN

Centre for Infectious Diseases and Microbiology Laboratory Services, ICPMR - Pathology West, Sydney Medical School, University of Sydney, and Westmead Hospital, Darcy Road, Westmead, NSW 2145, Australia

SUNWEN CHOU

Division of Infectious Diseases, Department of Medicine, Oregon Health & Science University, Portland OR 97239

JENS JØRGEN CHRISTENSEN

Department of Clinical Microbiology, Slagelse Hospital, Slagelse, Denmark

3/19/15 1:07 PM

CONTRIBUTORS

OANA CIOFU

G. SYBREN de HOOG

JÉRÔME CLAIN

PETER DEPLAZES

TOM COENYE

EDWARD P. DESMOND

Department of Immunology and Microbiology, Faculty of Health Sciences, University of Copenhagen, Copenhagen, Denmark UMR 216 IRD, Faculté de Pharmacie, Université Paris Descartes, 4 av de l’Observatoire, 75006 Paris, France Laboratory of Pharmaceutical Microbiology, Ghent University, B-9000 Gent, Belgium

GEORG CONRADS

Division of Oral Microbiology and Immunology, RWTH Aachen University Hospital, 52074 Aachen, Germany

PATRICIA S. CONVILLE

Microbiology Service, Department of Laboratory Medicine, Warren Grant Magnuson Clinical Center, National Institutes of Health, U.S. Department of Health and Human Services, Bethesda, MD 20892

PIET COOLS

CBS Fungal Biodiversity Centre, Institute of the Royal Netherlands Academy of Arts and Sciences, Uppsalalaan 8, 3584 CT, Utrecht, The Netherlands Institute of Parasitology, University of Zurich, Winterthurerstrasse 266a, CH-8057 Zurich, Switzerland Mycobacteriology Section, Microbial Diseases Laboratory, California Department of Public Health, Richmond, CA 94804

DANIEL J. DIEKEMA

Division of Infectious Diseases, Department of Internal Medicine, and Division of Medical Microbiology, Department of Pathology, University of Iowa College of Medicine, Iowa City, IA 52242

CHRISTOPHER D. DOERN

Department of Pathology, Virginia Commonwealth University Medical Center, Richmond, VA 23298

Laboratory for Bacteriology Research, Department Clinical Chemistry, Microbiology & Immunology, Faculty of Medicine & Health Sciences, University of Ghent, 3BlokA, De Pintelaan 185, UZ Gent, 9000 Gent, Belgium

GARY V. DOERN

SARAH D. COPSEY

Unité Mycologie Moléculaire, CNRS URA3012, Centre National de Référence des Mycoses Invasives et Antifongiques, Institut Pasteur, Paris, France

Public Health Wales, Anaerobe Reference Laboratory, University Hospital of Wales, Heath Park, Cardiff, CF14 4XW, United Kingdom

NATALIE A. COUNIHAN

School of Medicine, Deakin University, Waurn Ponds, 3218, Australia

DAVID L. COX

Emeritus, Department of Pathology, Carver College of Medicine, University of Iowa, Iowa City, IA 52242

FRANÇOISE DROMER

J. STEPHEN DUMLER

Departments of Pathology and Microbiology & Immunology, University of Maryland School of Medicine, 322D Health Science Facility–I, 685 West Baltimore St., Baltimore, MD 21201

Treponema Immunobiology and Syphilis Serology Activities, Division of STD Prevention, Laboratory Reference and Research Branch, Centers for Disease Control and Prevention, Atlanta, GA 30333

JAMES J. DUNN

FRANCIS E. G. COX

MARCELA ECHAVARRIA

BART J. CURRIE

PAUL H. EDELSTEIN

MELANIE T. CUSHION

JOHANNES ELIAS

Department of Disease Control, London School of Hygiene and Tropical Medicine, London WC1E 7HT, United Kingdom Menzies School of Health Research and Northern Territory Medical Program, Royal Darwin Hospital, Darwin, Northern Territory 0811, Australia University of Cincinnati College of Medicine, Cincinnati, OH 45267-0555, and Cincinnati Veterans Affairs Medical Center, Cincinnati, OH 45220

INGER K. DAMON

Centers for Disease Control and Prevention, Mailstop G-06, 1600 Clifton Rd. NE, Atlanta, GA 30030

MCM11_FM.indd 15

n  xv

Department of Pathology, Texas Children’s Hospital, and Department of Pathology and Immunology, Baylor College of Medicine, Houston, TX 77030 Clinical Virology Unit, CEMIC University Hospital, Galvan 4102 (1431 FWO) Buenos Aires, Argentina Department of Pathology and Laboratory Medicine, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA 19104-4283 Institute for Hygiene and Microbiology, University of Würzburg, 97080 Würzburg, Germany

HERMES ESCALANTE

Department of Microbiology and Parasitology, School of Biological Sciences, Universidad Nacional de Trujillo, Jr. Bolognesi 334, Trujillo, Peru

3/19/15 1:07 PM

xvi

n  CONTRIBUTORS

ANDREAS ESSIG

Institute of Medical Microbiology and Hygiene, University Ulm, Albert-Einstein Allee 23, D-89081 Ulm, Germany

RICHARD R. FACKLAM

Streptococcus Laboratory, Respiratory Diseases Branch, Division of Bacterial Diseases, Centers for Disease Control and Prevention, Mail Stop C0-2, Atlanta, GA 30333

FERRIC C. FANG

Departments of Laboratory Medicine and Microbiology, University of Washington School of Medicine, Seattle, WA 98195

J. J. FARMER, III

Silver Hill Associates, 1781 Silver Hill Road, Stone Mountain, GA 30087-2212

PATRICIA I. FIELDS

Enteric Diseases Laboratory Branch, Centers for Disease Control and Prevention, Atlanta, GA 30333

SYDNEY M. FINEGOLD

NAOMI J. GADSBY

Specialist Virology Centre, Department of Laboratory Medicine, Royal Infirmary of Edinburgh, 51 Little France Crescent, Edinburgh, EH16 4SA, United Kingdom

HECTOR H. GARCIA

Center for Global Health, Department of Microbiology, Universidad Peruana Cayetano Heredia, & Cysticercosis Unit, Instituto Nacional de Ciencias Neurologicas, H. Delgado 430, SMP, Lima 31, Peru

LYNNE S. GARCIA

LSG & Associates, 512 12th Street, Santa Monica, CA 90402

DEA GARCIA-HERMOSO

Unité Mycologie Moléculaire, CNRS URA3012, Centre National de Référence des Mycoses Invasives et Antifongiques, Institut Pasteur, Paris, France

BARBARA C. GÄRTNER

Virology, Institute for Microbiology, University of the Saarland, D-66421 Homburg/Saar, Germany

Infectious Diseases Section, VA Medical Center, and Department of Medicine and Department of Microbiology, Immunology, and Molecular Genetics, University of California at Los Angeles, Los Angeles, CA 90073

CHARLOTTE A. GAYDOS

COLLETTE FITZGERALD

WALTER GEIßDÖRFER

National Campylobacter and Helicobacter Reference Laboratory, Enteric Diseases Laboratory Branch, Centers for Disease Control and Prevention, Atlanta, GA 30329

MICHAEL S. FORMAN

Division of Medical Microbiology, Department of Pathology, The Johns Hopkins Hospital, 600 North Wolfe Street, Meyer B1-193, Baltimore, MD 21287-7093

STEPHEN J. FORSYTHE

Nottingham Trent University, Nottingham, NG11 8NS, United Kingdom

RICHARD FRANKA

Poxvirus and Rabies Branch, Division of High Consequence Pathogens and Pathology, Centers for Disease Control and Prevention, Atlanta, GA 30333

RENO FREI

Universitätsspital Basel, Klinische Mikrobiologie, Petersgraben 4, CH-4031 Basel, Switzerland

MATTHIAS FROSCH

Institute for Hygiene and Microbiology, University of Würzburg, 97080 Würzburg, Germany

CHARLES F. FULHORST

Department of Pathology, The University of Texas Medical Branch at Galveston, Galveston, TX 77555-0609

GUIDO FUNKE

Risch Laboratories Group, Landstrasse 157, FL-9494 Schaan, Principality of Liechtenstein

MCM11_FM.indd 16

Johns Hopkins University, Division of Infectious Diseases, Department of Medicine, 530 Rangos Building, 855 N. Wolfe St., Baltimore, MD 21205 Mikrobiologisches Institut - Klinische Mikrobiologie, Immunologie und Hygiene, Friedrich-Alexander-Universität Erlangen-Nürnberg and Universitätsklinikum Erlangen, 91054 Erlangen, Germany

PETER GERNER-SMIDT

National Center for Emerging and Zoonotic Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, GA 30333

MAHMOUD A. GHANNOUM

Center for Medical Mycology, Department of Dermatology, Case Western Reserve University, Cleveland, OH 44106

CHRISTINE C. GINOCCHIO

Hofstra North Shore-LIJ School of Medicine, 500 Hofstra University, Hempstead, NY 11549, and bioMérieux, 100 Rodolphe St., Durham, NC 27712

LORI M. GLADNEY

IHRC Inc. contractor at the Enteric Diseases Laboratory Branch, Division of Foodborne, Waterborne and Environmental Diseases, Centers for Disease Control and Prevention, Atlanta, GA 30333

MARKUS GLATZEL

Institute of Neuropathology, University Medical Center HamburgEppendorf, Martinistraße 52, D-20246 Hamburg, Germany

BEATRIZ L. GÓMEZ

School of Medicine and Health Sciences, Universidad Rosario, Bogota, and Medical and Experimental Mycology Unit, Corporación para Investigaciones Biológicas (CIB), Medellín, Colombia

3/19/15 1:07 PM

CONTRIBUTORS n  xvii

STEPHEN R. GRAVES

AIMEE C. HODOWANEC

PATTI E. GRAVITT

NIELS HØIBY

Australian Rickettsial Reference Laboratory, Geelong, Victoria, Australia Department of Pathology, University of New Mexico Health Sciences Center, MSC08 4640, 1 University of New Mexico, Albuquerque, NM 87131-0001

JOSEP GUARRO

Medical School & IISPV, University Rovira i Virgili, Sant Llorenç, 21, 43201 Reus, Spain

VAL HALL

Public Health Wales, Anaerobe Reference Laboratory, University Hospital of Wales, Heath Park, Cardiff, CF14 4XW, United Kingdom

CATHLEEN A. HANLON

Poxvirus and Rabies Branch, Division of High Consequence Pathogens and Pathology, Centers for Disease Control and Prevention, Atlanta, GA 30333

PATRICIA HARRIS

Harris Consulting, Henderson, NV 89044

RANDALL T. HAYDEN

Department of Pathology, St. Jude Children’s Research Hospital, Memphis, TN 38105-3678

KEVIN C. HAZEN

Clinical Microbiology Laboratory, Department of Pathology, Duke University Health System, Durham, NC 27710

DAVID W. HECHT

Department of Medicine, Loyola University Medical Center, Maywood, IL 60153

PEERA HEMARAJATA

Clinical Microbiology Laboratory, Pathology and Laboratory Medicine, University of California Los Angeles, Los Angeles, CA 90049

WALID HENEINE

Laboratory Branch, Division of HIV/AIDS Prevention, National Center for HIV/AIDS, Viral Hepatitis, STD, and TB Prevention, Centers for Disease Control and Prevention, 1600 Clifton Rd., MS G-45, Atlanta, GA 30329

SARAH K. HIGHLANDER

Department of Molecular Virology & Microbiology, Baylor College of Medicine, Houston, TX 77030

JANET A. HINDLER

Department of Pathology and Laboratory Medicine, University of California, Los Angeles, Los Angeles, CA 90095

RICHARD L. HODINKA

Department of Biomedical Sciences, University of South Carolina School of Medicine Greenville and Greenville Health System, Greenville, SC 29605

MCM11_FM.indd 17

Department of Medicine, Section of Infectious Diseases, Rush University Medical Center, Chicago, IL 60612 Department of Clinical Microbiology, Rigshospitalet, and Department of Immunology and Microbiology, Faculty of Health Sciences, University of Copenhagen, Copenhagen, Denmark

MARTIN HOLFELDER

Department of Microbiology, Limbach Laboratory, D-69120 Heidelberg, Germany

AMY J. HORNEMAN

Pathology and Laboratory Service, Microbiology and Molecular Diagnostics, VA Maryland Health Care System, Baltimore, MD 21201-1524

REBECCA T. HORVAT

Department of Pathology and Laboratory Medicine, University of Kansas School of Medicine, 3901 Rainbow Blvd., Mail Stop 3045, Kansas City, KS 66160

SUSAN A. HOWELL

Mycology, St. Johns Institute of Dermatology, Viapath, London, SE1 7EH, United Kingdom

DIANA D. HUANG

Department of Immunology/Microbiology, Rush University Medical Center, 1653 W. Congress Parkway, Chicago, IL 60612

LAURA HUGHES

Centers for Disease Control and Prevention, Mailstop G-06, 1600 Clifton Rd. NE, Atlanta, GA 30030

ROMNEY M. HUMPHRIES

Department of Pathology and Laboratory Medicine, University of California, Los Angeles, Los Angeles, CA 90095

ELIZABETH HUNSPERGER

Immunodiagnostics, Development and Research Laboratory, CDC, National Centers for Emerging and Zoonotic Infectious Diseases, Division of Vector-Borne Diseases, Dengue Branch, 1324 Calle Canada, San Juan, Puerto Rico 00920

JOSEPH P. ICENOGLE

Measles, Mumps, Rubella and Herpesviruses Laboratory Branch, Mail Stop C-22, Centers for Disease Control and Prevention, 1600 Clifton Road, NE, Atlanta, GA 30333

NANCY C. ISHAM

Case Western Reserve University, Cleveland, OH 44106

KEITH R. JEROME

Department of Laboratory Medicine, University of Washington, and Vaccine and Infectious Disease Division, Fred Hutchinson Cancer Research Center, 1100 Fairview Ave N., E5-110, Seattle, WA 98109

3/19/15 1:07 PM

xviii

n  CONTRIBUTORS

JUAN A. JIMENEZ

Department of Microbiology, Universidad Peruana Cayetano Heredia, H. Delgado 430, SMP, Lima 31, Peru

ELIZABETH M. JOHNSON

MARIE LOUISE LANDRY

Departments of Laboratory Medicine and Internal Medicine, Yale University School of Medicine, P.O. Box 208035, New Haven, CT 06520-8035

Mycology Reference Laboratory, Public Health England, South West Laboratory, Myrtle Road, Kingsdown, Bristol BS2 8EL, United Kingdom

MARK T. LaROCCO

JEFFREY L. JONES

Gastrointestinal Bacteriology Reference Unit, Public Health England, 61 Colindale Avenue, London, NW9 5EQ, United Kingdom

Parasitic Diseases Branch, Division of Parasitic Diseases, Center for Global Health, Centers for Disease Control and Prevention, 1600 Clifton Road NE, MS F-22, Atlanta, GA 30329

MALCOLM K. JONES

School of Veterinary Sciences, The University of Queensland, Brisbane, QLD 4072, Australia

JAMES H. JORGENSEN

Emeritus, Department of Pathology, University of Texas Health Science Center, 7703 Floyd Curl Dr., San Antonio, TX 78229-3900

PETER KÄMPFER

Institut für Angewandte Mikrobiologie Justus-LiebigUniversität Giessen, D-35392 Giessen, Germany

Erie, PA 16506

ANDY J. LAWSON

JACQUES LE BRAS

UMR 216 IRD, Faculté de Pharmacie, Université Paris Descartes, 4 av de l’Observatoire, 75006 Paris, France

AMY L. LEBER

Nationwide Children’s Hospital, Clinical Microbiology and Immunoserology, Department of Laboratory Medicine, 700 Children’s Drive, Bldg C, Rm 1868, Columbus, OH 43205

NATHAN A. LEDEBOER

Department of Pathology, Medical College of Wisconsin, Milwaukee, WI 53226

KARIN LEDER

Department of Microbiology and Immunology, University of Maryland School of Medicine, Baltimore, MD 21201

Infectious Disease Epidemiology Unit, School of Epidemiology and Preventive Medicine, Monash University, and Victorian Infectious Diseases Service, Royal Melbourne Hospital at the Doherty Institute for Infection and Immunity, Victoria, Australia

JAMES A. KARLOWSKY

ELLIOT J. LEFKOWITZ

JAMES B. KAPER

Diagnostic Services Manitoba and Department of Medical Microbiology, University of Manitoba, Winnipeg, Manitoba, R2H 2A6, Canada

JENNIFER KEISER

Department of Medical Parasitology and Infection Biology, Swiss Tropical and Public Health Institute, P.O. Box, CH-4002 Basel, Switzerland

NANCY KELLER

Department of Medical Microbiology & Immunology, University of Wisconsin, Madison, WI 53705

EIJA KÖNÖNEN

Institute of Dentistry, University of Turku, 20520 Turku, Finland

THOMAS G. KSIAZEK

Galveston National Laboratory, Department of Pathology, University of Texas Medical Branch, Galveston, TX 77555

JAIME LABARCA

Dept. Internal Medicine, Pontificia Universidad de Chile, Lira 63, Santiago, Chile

DARYL M. LAMSON

Laboratory of Viral Diseases, Wadsworth Center, Albany, NY 12201

MCM11_FM.indd 18

Department of Microbiology, The University of Alabama at Birmingham, Birmingham, AL 35294

DIANE S. LELAND

Department of Pathology and Laboratory Medicine, Indiana University School of Medicine, IU Health Pathology Laboratory Building, Room 6027F, 350 W. 11th Street, Indianapolis, IN 46202

PAUL N. LEVETT

Saskatchewan Disease Control Laboratory, Regina, Saskatchewan, Canada

JAMES S. LEWIS, II

Departments of Pharmacy and Infectious Diseases, Oregon Health and Science University, Portland, OR 97239

BRANDI M. LIMBAGO

Centers for Disease Control and Prevention, Atlanta, GA 30329

SHOU-YEAN GRACE LIN

Mycobacteriology Section, Microbial Diseases Laboratory, California Department of Public Health, Richmond, CA 94804

DAVID S. LINDSAY

Center for Molecular Medicine and Infectious Diseases, Department of Biomedical Sciences and Pathobiology, VirginiaMaryland Regional College of Veterinary Medicine, Virginia Tech, 1410 Prices Fork Road, Blacksburg, VA 24061-0342

3/19/15 1:07 PM

CONTRIBUTORS n  xix

MARK D. LINDSLEY

KARIN L. McGOWAN

STEPHEN E. LINDSTROM

DONALD P. McMANUS

Mycotic Diseases Branch, Centers for Disease Control and Prevention, 1600 Clifton Road, Mailstop G-11, Atlanta, GA 30333 Influenza Branch, MS-G16, Division of Viral and Rickettsial Diseases, National Center for Infectious Diseases, Centers for Disease Control and Prevention, 1600 Clifton Road, Atlanta, GA 30333

Emeritus Professor, Department of Pathology and Laboratory Medicine, Perelman School of Medicine, University of Pennsylvania, Philadelphia, PA 19104 QIMR Berghofer Medical Research Institute, 300 Herston Road, Herston, Queensland 4006, Australia

LEONEL MENDOZA

Ochsner Medical Center, Dept. of Pathology, 1514 Jefferson Highway, New Orleans, LA 70121

Biomedical Laboratory Diagnostics, Microbiology and Molecular Genetics, Michigan State University, North Kedzie Hall, 354 Farm Lane Room N326, East Lansing, MI 48824-1031

JOHN J. LiPUMA

WIELAND MEYER

ANDREA J. LINSCOTT

Department of Pediatrics and Communicable Diseases, University of Michigan Medical School, Ann Arbor, MI 48019

SHAWN R. LOCKHART

National Center for Emerging and Zoonotic Infectious Diseases, Centers for Disease Control and Prevention, 1600 Clifton Road NE, Mailstop G-11, Atlanta, GA 30333

Molecular Mycology Research Laboratory, Ctr. for Infectious Diseases & Microbiology, Sydney Medical School, and Marie Bashir Institute for Infectious Diseases and Biosecurity, University of Sydney; and Westmead Hospital, Westmead, NSW 2145, Australia

RHODA ASHLEY MORROW

University of Texas Medical Branch, 301 University Blvd., Galveston, TX 77555-0740

Department of Laboratory Medicine, University of Washington, and Vaccine and Infectious Disease Division, Fred Hutchinson Cancer Research Center, 1100 Fairview Ave N., E5-110, Seattle, WA 98109

OLIVIER LORTHOLARY

ANNETTE MOTER

MICHAEL LOEFFELHOLZ

Unité Mycologie Moléculaire, CNRS URA3012, Centre National de Référence des Mycoses Invasives et Antifongiques, Institut Pasteur, Paris, France

XIAOYAN LU

Biofilmcenter, German Heart Institute Berlin and CharitéUniversitätsmedizin Berlin, 12203 Berlin, Germany

IONA MUNJAL

Division of Viral Diseases, Centers for Disease Control and Prevention, Atlanta, GA 30333

Department of Pediatrics, Division of Infectious Diseases, The Children’s Hospital at Montefiore, and Albert Einstein College of Medicine, Bronx, NY 10467

CHRISTIAN LÜCK

IRVING NACHAMKIN

Institut für Medizinische Mikrobiologie und Hygiene, Technische Universität Dresden, 01307 Dresden, Germany

NELL S. LURAIN

Department of Immunology/Microbiology, Rush University Medical Center, Chicago, IL 60612

DUNCAN MacCANNELL

National Center for Emerging and Zoonotic Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, GA 30333

ROBERT F. MASSUNG

Department of Pathology and Laboratory Medicine, Perelman School of Medicine, University of Pennsylvania, 7th Floor Founders Building, 3400 Spruce Street, Philadelphia, PA 19104-4283

ELISABETH NAGY

Institute of Clinical Microbiology, University of Szeged, 6725 Szeged, Hungary

JAMES P. NATARO

Department of Pediatrics, University of Virginia School of Medicine, Charlottesville, VA 22908

Division of Vector-Borne Diseases, National Center for Emerging and Zoonotic Infectious Diseases, Centers for Disease Control and Prevention, 1600 Clifton Road, N.E., MS G-13, Atlanta, GA 30333

RONALD C. NEAFIE

ALEXANDER MATHIS

Laboratory of Bacterial Genetics, National Institute of Public Health, Srobarova 48, 10042 Prague, Czech Republic

Institute of Parasitology, University of Zurich, Winterthurerstrasse 266a, CH-8057 Zurich, Switzerland

JAMES B. McAULEY

Rush University Medical Center, 1653 W. Congress Parkway, Chicago, IL 60612

MCM11_FM.indd 19

Parasitic Infections Branch, Armed Forces Institute of Pathology, 6825 16th Street NW, Washington, DC 20306

ALEXANDR NEMEC

STUART T. NICHOL

Viral Special Pathogens Branch, MS G-14, Centers for Disease Control and Prevention, 1600 Clifton Rd., NE, Atlanta, GA 30333

3/19/15 1:07 PM

xx

n  CONTRIBUTORS

MICHAEL A. NOBLE

Department of Pathology and Laboratory Medicine, University of British Columbia, Vancouver BC V6T 2B5, Canada

FREDERICK S. NOLTE

Department of Pathology and Laboratory Medicine, Medical University of South Carolina, Charleston, SC 29425

SUSAN NOVAK-WEEKLEY

Microbiology, Immunoserology and Molecular Infectious Disease, SCMPG Regional Reference Laboratories, 11668 Sherman Way, North Hollywood, CA 91605

THOMAS B. NUTMAN

Laboratory of Parasitic Diseases, National Institute of Allergy and Infectious Diseases, National Institutes of Health, Bethesda, MD 20892

KERRY O’DONNELL

SHARON J. PEACOCK

Department of Medicine, University of Cambridge, Addenbrooke’s Hospital, Cambridge, CB2 0QQ, United Kingdom

PHILIP E. PELLETT

Department of Immunology and Microbiology, Wayne State University School of Medicine, Scott Hall Room 6225, 540 East Canfield Ave., Detroit, MI 48201

DAVID S. PERLIN

Public Health Research Institute, New Jersey Medical School, Rutgers Biomedical and Health Sciences, Newark, NJ 07103

JEANNINE M. PETERSEN

Bacterial Diseases Branch, Division of Vector-Borne Diseases, Centers for Disease Control and Prevention, Fort Collins, CO 80521

Bacterial Foodborne Pathogens and Mycology, National Center for Agricultural Utilization Research, ARS-USDA, 1815 N. University St., Peoria, IL 61604-3902

JOSEPH F. PETROSINO

VICTORIA A. OLSON

CATHY A. PETTI

Centers for Disease Control and Prevention, Mailstop G-06, 1600 Clifton Rd. NE, Atlanta, GA 30030

LILLIAN A. ORCIARI

Poxvirus and Rabies Branch, Division of High Consequence Pathogens and Pathology, Centers for Disease Control and Prevention, Atlanta, GA 30333

BELINDA OSTROWSKY

Department of Medicine, Division of Infectious Disease, Montefiore Medical Center, and Albert Einstein College of Medicine, Bronx, NY 10467

S. MICHELE OWEN

Laboratory Branch, Division of HIV/AIDS Prevention, National Center for HIV/AIDS, Viral Hepatitis, STD, and TB Prevention, Centers for Disease Control and Prevention, 1600 Clifton Rd., MS G-45, Atlanta, GA 30329

GRAEME P. PALTRIDGE

Canterbury Health Laboratories, Microbiology Unit, P.O. Box 151, Hagley Ave. and Tuam St., Christchurch, 8011, New Zealand

XIAOLI PANG

Provincial Laboratory for Public Health, Walter Mackenzie Health Sciences Centre, University of Alberta Hospital, 8440 - 112 St., Edmonton, AB, T6G 2J2, Canada

JEAN B. PATEL

Office of Antimicrobial Resistance, Division of Healthcare Quality Promotion, Centers for Disease Control and Prevention, MS C-12, 1600 Clifton Rd. NE, Atlanta, GA 30333

ROBIN PATEL

Mayo Clinic, Department of Laboratory Medicine and Pathology, Division of Clinical Microbiology and Department of Medicine, Division of Infectious Diseases, Rochester, MN 55905

MCM11_FM.indd 20

Department of Molecular Virology & Microbiology, Baylor College of Medicine, Houston, TX 77030 HealthSpring Global Inc., Bradenton, FL 34209

MICHAEL A. PFALLER

T2 Biosystems, Lexington, MA 02421, and Professor Emeritus, University of Iowa College of Medicine and College of Public Health, Iowa City, IA 52242

GABY E. PFYFFER

Department of Medical Microbiology, Center for Laboratory Medicine, Luzerner Kantonsspital, 6000 Luzern 16, Switzerland

ALLAN PILLAY

Molecular Diagnostics & Typing Laboratory, Division of STD Prevention, Laboratory Reference and Research Branch, Centers for Disease Control and Prevention, Atlanta, GA 30333

JOHANN PITOUT

Calgary Laboratory Services, University of Calgary, Calgary, Alberta, Canada

JUTTA PREIKSAITIS

University of Alberta, 2B2.08 WMC, 8440 112 Street, Edmonton, AB T6G 2B7, Canada

BOBBI S. PRITT

Mayo Clinic, Department of Laboratory Medicine and Pathology, Division of Clinical Microbiology, 200 1st Street SW, Rochester, MN 55905

GARY W. PROCOP

Department of Laboratory Medicine, Pathology and Laboratory Medicine Institute, Cleveland Clinic, Cleveland, OH 44195

ELISABETH PUCHHAMMER-STÖCKL

Department of Virology, Medical University of Vienna, Kinderspitalgasse 15, A-1095 Vienna, Austria

3/19/15 1:07 PM

CONTRIBUTORS n  xxi

JUSTIN D. RADOLF

Departments of Medicine, Pediatrics, Immunology, Molecular Biology and Biophysics, and Genetics and Developmental Biology, University of Connecticut Health Center, Farmington, CT 06030

L. BARTH RELLER

Departments of Pathology and Medicine, Duke University School of Medicine, Durham, NC 27710

MEGAN E. RELLER

MPH Division of Medical Microbiology, Department of Pathology, The Johns Hopkins University School of Medicine, 720 Rutland Avenue, Ross 628, Baltimore, MD 21205

ELVIRA RICHTER

Forschungszentrum Borstel, Leibniz-Zentrum für Medizin und Biowissenschaften, Nationales Referenzzentrum für Mykobakterien, Parkallee 18, 23845 Borstel, Germany

SANDRA S. RICHTER

Department of Laboratory Medicine, Cleveland Clinic, 9500 Euclid Avenue, LL1-2, Cleveland, OH 44195

MARION RIFFELMANN

Labor:Medizin Krefeld MVZ, D-47805 Krefeld, Germany

CHRISTINE ROBINSON

Department of Pathology, Children’s Hospital Colorado, Aurora, CO 80045

PIERRE E. ROLLIN

Viral Special Pathogens Branch, MS G-14, Centers for Disease Control and Prevention, 1600 Clifton Rd., NE, Atlanta, GA 30333

JOSÉ R. ROMERO

Pediatric Infectious Diseases Section, University of Arkansas for Medical Sciences and Arkansas Children’s Hospital, Little Rock, AR 72202-3591

PAUL A. ROTA

National Center for Immunization and Respiratory Diseases, Centers for Disease Control and Prevention, Atlanta, GA 30333

KATHRYN L. RUOFF

O’Toole Lab, Dept. of Microbiology and Immunology, Geisel School of Medicine at Dartmouth, Hanover, NH 03755

NORBERT RYAN

Bacteriology Laboratory, Victorian Infectious Diseases Reference Laboratory, Melbourne, Victoria 3000, Australia

MARTIN E. SCHRIEFER

Bacterial Diseases Branch, Division of Vector-Borne Diseases, Centers for Disease Control and Prevention, Fort Collins, CO 80521

AUDREY N. SCHUETZ

Department of Pathology and Laboratory Medicine, Weill Cornell Medical College, New York, NY 10065

MCM11_FM.indd 21

DIANA G. SCORPIO

Veterinary Medicine, Ross University School of Veterinary Medicine, St. Kitts, West Indies

W. EVAN SECOR

Division of Parasitic Diseases and Malaria, Centers for Disease Control and Prevention, 1600 Clifton Rd, N.E., MS D-65, Atlanta, GA 30329

ARLENE C. SEÑA

Department of Medicine, Division of Infectious Diseases, University of North Carolina at Chapel Hill, Chapel Hill, NC 27599

ROBERT W. SHAFER

Division of Infectious Diseases, Lane Building L143, Stanford University, Stanford, CA 94305

SUSAN E. SHARP

Kaiser Permanente, Department of Pathology, 13705 NE Airport Way, Portland, OR 97230

ROSEMARY C. SHE

Department of Pathology, Keck School of Medicine of USC, Los Angeles, CA 90033

HARSHA SHEOREY

Department of Microbiology, St Vincent’s Hospital Melbourne, Fitzroy, Victoria 3065, Australia

PATRICIA LYNN SHEWMAKER

Streptococcus Laboratory, Respiratory Diseases Branch, Division of Bacterial Diseases, Centers for Disease Control and Prevention, Mail Stop C0-2, Atlanta, GA 30333

ROBYN Y. SHIMIZU

UCLA Health, Department of Pathology and Laboratory Medicine, 11633 San Vicente Blvd., Rear Building, Los Angeles, CA 90049

PATRICIA J. SIMNER

Division of Clinical Microbiology, Mayo Clinic, Rochester, MN 55905

KAMALJIT SINGH

Rush University Medical Center, Academic Facility, Suite 140, 600 S. Paulina, Chicago, IL 60612

ROBERT L. SKOV

Statens Serum Institut, Department of Microbiological Surveillance and Research, Copenhagen 2300, Denmark

JENNIFER S. SMITH

Department of Epidemiology, University of North Carolina, Chapel Hill, NC 27599

JAMES W. SNYDER

Department of Pathology and Laboratory Medicine, University of Louisville School of Medicine, 530 S. Jackson St., Louisville, KY 40202

YULI SONG

Mason Business Center, Proctor & Gamble, Mason, OH 45040

3/19/15 1:07 PM

xxii

n  CONTRIBUTORS

TANIA C. SORRELL

Centre for Infectious Diseases and Microbiology, Sydney Medical School, University of Sydney; Marie Bashir Institute for Infectious Diseases and Biosecurity, University of Sydney; and Westmead Hospital, Westmead, NSW 2145, Australia

BARBARA SPELLERBERG

Institute of Medical Microbiology and Hygiene, University of Ulm, Albert Einstein Allee 11, 89081 Ulm, Germany

JENNIFER K. SPINLER

Department of Pathology, Baylor College of Medicine, and Department of Pathology, Texas Children’s Hospital, Houston, TX 77030

KATHLEEN A. STELLRECHT

Microbiology, Department of Pathology and Laboratory Medicine, Albany Medical College and Albany Medical Center Hospital, Albany Medical Center, Albany, NY 12208

STEFFEN STENGER

Universitätsklinikum Ulm, Institut für Med. Mikrobiologie und Hygiene, Albert Einstein Allee 11 (M23/4204), D-89081 Ulm, Germany

DENNIS L. STEVENS

Veterans Affairs Medical Center, Boise, ID 83702, and University of Washington School of Medicine, Seattle, WA 98195

NANCY A. STROCKBINE

Enteric Diseases Laboratory Branch, Centers for Disease Control and Prevention, Atlanta, GA 30333

RICHARD C. SUMMERBELL

Sporometrics Inc., and Dalla Lana School of Public Health, University of Toronto, Toronto, Ontario, Canada

DEANNA A. SUTTON

Fungus Testing Laboratory, Department of Pathology, University of Texas Health Science Center at San Antonio, San Antonio, TX 78229-3900

JANA M. SWENSON

Centers for Disease Control and Prevention, Atlanta, GA 30333

WILLIAM M. SWITZER

Laboratory Branch, Division of HIV/AIDS Prevention, National Center for HIV/AIDS, Viral Hepatitis, STD, and TB Prevention, Centers for Disease Control and Prevention, 1600 Clifton Rd., MS G-45, Atlanta, GA 30329

YI-WEI TANG

Department of Laboratory Medicine, Memorial SloanKettering Cancer Center; Department of Pathology and Laboratory Medicine, Weill Medical College of Cornell University, New York, NY 10065

MCM11_FM.indd 22

CHERYL L. TARR Enteric Diseases Laboratory Branch, Centers for Disease Control and Prevention, Atlanta, GA 30333

RYAN TAYLOR

Department of Medicine, Division of Gastroenterology, Hepatology and Motility, University of Kansas School of Medicine, 3901 Rainbow Blvd., Mail Stop 3045, 4035 Wescoe Pavilion, Kansas City, KS 66160

DAVID TAYLOR-ROBINSON

Section of Infectious Diseases, Wright-Fleming Institute, Faculty of Medicine, Imperial College London, St Mary’s Campus, Norfolk Place, London W2 1PG, United Kingdom

LUCIA MARTINS TEIXEIRA

Instituto de Microbiologia, Universidade Federal do Rio de Janeiro, Rio de Janeiro, RJ 21941-902, Brazil

SAM R. TELFORD III

Cummings School of Veterinary Medicine, Tufts University, 200 Westboro Road, North Grafton, MA 01536

KATE E. TEMPLETON

Specialist Virology Centre, Department of Laboratory Medicine, Royal Infirmary of Edinburgh, 51 Little France Crescent, Edinburgh, EH16 4SA, United Kingdom

ELITZA S. THEEL

Department of Laboratory Medicine and Pathology, Division of Clinical Microbiology, Mayo Clinic, Rochester, MN 55901

KENNETH D. THOMPSON

Department of Pathology, The University of Chicago Medical Center, Chicago, IL 60637

GEORGE R. THOMPSON, III

Department of Medical Microbiology and Immunology, Department of Medicine, Division of Infectious Diseases, University of California - Davis, Davis, CA 95616 (retired)

KURT THROCKMORTON

Genetics Department, University of Wisconsin, 425 Henry Mall, Madison, WI 53704

GRAHAM TIPPLES

Provincial Laboratory for Public Health, Alberta Health Services, and Department of Medical Microbiology and Immunology, University of Alberta, Edmonton, AB T6G 2J2, Canada

EIJA TREES

National Center for Emerging and Zoonotic Infectious Diseases, Centers for Disease Control and Prevention, Atlanta, GA 30333

CHRISTINE Y. TURENNE

Saskatchewan Disease Control Laboratory, Saskatchewan Ministry of Health, Regina, Saskatchewan S4S 0A4, Canada

3/19/15 1:07 PM

CONTRIBUTORS n  xxiii JOHN D. TURNIDGE Departments of Pathology, Paediatrics and Molecular and Biomedical Sciences, University of Adelaide, and SA Pathology, Women’s and Children’s Hospital, North Adelaide 5006, South Australia

DAVID H. WALKER Department of Pathology, Center for Biodefense and Emerging Infectious Diseases, University of Texas Medical Branch, Galveston, TX 77555-0609

ALEXANDRA VALSAMAKIS

University of Texas Health Science Center at Tyler, Department of Microbiology, 11937 U.S. Highway 271, Tyler, TX 75708

Division of Medical Microbiology, Department of Pathology, The Johns Hopkins Hospital, 600 North Wolfe Street, Meyer B1-193, Baltimore, MD 21287-7093

WENDY W. J. van de SANDE

Erasmus MC, Department of Medical Microbiology & Infectious Diseases, s-Gravendijkwal 230, 3015 CE Rotterdam, The Netherlands

GERALD van HORN

Microbiology Laboratory, Winthrop University Hospital, Mineola, NY 11501

PETER A. R. VANDAMME

Laboratorium voor Microbiologie, Faculteit Wetenschappen, Universiteit Gent, Ledeganckstraat 35, B-9000 Gent, Belgium

MARIO VANEECHOUTTE

Laboratory for Bacteriology Research, Department of Clinical Chemistry, Microbiology & Immunology, Faculty of Medicine & Health Sciences, University of Ghent, 3BlokA, De Pintelaan 185, UZ Gent, 9000 Gent, Belgium

JAMES VERSALOVIC

Department of Pathology & Immunology and Department of Molecular Virology & Microbiology, Baylor College of Medicine, and Department of Pathology, Texas Children’s Hospital, Houston, TX 77030

RAQUEL VILELA

Biomedical Laboratory Diagnostics, Michigan State University, North Kedzie Hall, 354 Farm Lane Room N322, East Lansing, MI 48824-1031

GOVINDA S. VISVESVARA

RICHARD J.WALLACE, JR.

DAVID W. WARNOCK

Faculty of Medical and Human Sciences, University of Manchester, Manchester, United Kingdom

GEORGES WAUTERS

Emeritus, Microbiologie, Université Catholique de Louvain, Brussels, Belgium

RAINER WEBER

Division of Infectious Diseases and Hospital Epidemiology, University Hospital and University of Zurich, Raemistrasse 100, CH-8091 Zurich, Switzerland

MELVIN P. WEINSTEIN

Departments of Medicine and Pathology & Laboratory Medicine, Rutgers Robert Wood Johnson Medical School, New Brunswick, NJ 08903

LOUIS M. WEISS

Albert Einstein College of Medicine, 1300 Morris Park Avenue, Room 504 Forchheimer Building, Bronx, NY 10461

PETER F. WELLER

Harvard Medical School, Division of Infectious Diseases, and Allergy and Inflammation Division, Beth Israel Deaconess Medical Center, Harvard School of Public Health, Boston, MA 02115

NELE WELLINGHAUSEN

Department of Medical Microbiology, Laboratories Ravensburg Dr. Gärtner, D-88212 Ravensburg, Germany

CONSTANZE WENDT

MVZ Labor Limbach, 691260 Heidelberg, Germany

Centers for Disease Control & Prevention, Division of Foodborne, Waterborne & Environmental Diseases, Waterborne Diseases Prevention Branch, 1600 Clifton Road, Atlanta, GA 30333 (retired)

NANCY L.WENGENACK

ULRICH VOGEL

Universitätsspital Basel, Abteilung für Spitalhygiene, Petersgraben 4, CH-4031 Basel, Switzerland

Institute for Hygiene and Microbiology, University of Würzburg, 97080 Würzburg, Germany

CHRISTOF von EIFF

University Hospital Münster, Institute of Medical Microbiology, D-48149 Münster, and Pfizer Pharma GmbH, 10785 Berlin, Germany

KEN B. WAITES

Department of Pathology, University of Alabama at Birmingham, WP 230, 619 South 19th Street, Birmingham, AL 35249

Division of Clinical Microbiology, Mayo Clinic, Rochester, MN 55905

ANDREAS F. WIDMER

DANNY L. WIEDBRAUK

Virology and Molecular Biology Department, Warde Medical Laboratory, 300 W. Textile Rd., Ann Arbor, MI 48108

MICHAEL L. WILSON

Department of Pathology & Laboratory Services, Denver Health, Denver, CO 80204, and Department of Pathology, University of Colorado School of Medicine, Aurora, CO 80045

CARL-HEINZ WIRSING von KÖNIG

Labor:Medizin Krefeld MVZ, D-47805 Krefeld, Germany

MCM11_FM.indd 23

3/19/15 1:07 PM

xxiv

n  CONTRIBUTORS

FRANK G. WITEBSKY

SHERIF ZAKI

GAIL L. WOODS

REINHARD ZBINDEN

Retired, Microbiology Service, Department of Laboratory Medicine, Warren G. Magnuson Clinical Center, National Institutes of Health, Bethesda, MD 20892 Department of Pathology and Laboratory Medicine, University of Arkansas for Medical Sciences, and Department of Pathology, Arkansas Children’s Hospital, Little Rock, AR 72205

LIHUA XIAO

Division of Foodborne, Waterborne and Environmental Diseases, Centers for Disease Control and Prevention, Mail Stop D66, 1600 Clifton Road, Atlanta, GA 30329-4018

Infectious Disease Pathology Branch, Centers for Disease Control and Prevention, 1600 Clifton Rd., NE, Atlanta, GA 30333 Institute of Medical Microbiology, University of Zurich, Gloriastrasse 32, CH 8006 Zurich, Switzerland

SEAN X. ZHANG

Microbiology Laboratory, The Johns Hopkins Hospital, 600 N Wolfe Street, Meyer B1-193, Baltimore, MD 21287-7093

Acknowledgment of Previous Contributors The Manual of Clinical Microbiology is by its nature a continuously revised work which refines and extends the contributions of previous editions. Since its first edition in 1970, many eminent scientists have contributed to this important reference work. The American Society for Microbiology and its Publications Board gratefully acknowledge the contributions of all of these generous authors over the life of this Manual.

MCM11_FM.indd 24

3/19/15 1:07 PM

Preface

The Manual of Clinical Microbiology (MCM) is the most authoritative reference text in the field of clinical microbiology. This edition of the Manual benefited from the talents of a team of 22 editors and almost 250 authors who were supported by a very capable production team at ASM Press. This, the 11th edition, is presented after the usual 4-year publication cycle following the 10th edition. All of the editorial team are proud members of the American Society for Microbiology and strong supporters of its book publishing arm, ASM Press. We have followed in the footsteps of previous authors and editors of the Manual and remain steadfastly committed to the utmost quality and timeliness that the MCM readership has come to expect. For the first time, we have had co-editors in chief of MCM. The length and scope of the Manual now require this division of labor to ensure thoroughness and timeliness of the editing process. We hope that readers of the Manual will recognize the commitment to excellence by everyone associated with its production. We represent only the fifth and sixth editors in chief in the 45-year history of the Manual. We are grateful for the example set by our predecessors and by the sage advice offered by recent editors in chief Patrick Murray and James Versalovic. We offer our deep appreciation to Ken April, the production editor at the outset of this edition, and to Ellie Tupper, who succeeded him and completed the editorial production process. This is only the second edition of the Manual to have a full-scale, searchable, Web-based HTML electronic edition. We hope that users of the Manual will find this

electronic alternative to the print version of MCM to be convenient and user friendly. It is likely that future editions of MCM will rely more heavily on the electronic format for delivery of the vast content of the Manual. This is a very dynamic era in clinical microbiology, with new technical tools (MALDI-TOF, ribosomal and total gene sequencing, and other molecular methods) that are profoundly influencing our approaches to organism detection and identification. The Manual continues to include classic microbiological techniques such as microscopy and culture as a foundation in addition to the newer methods cited above. Some organisms have become prominent causes of disease recently, e.g., Ebola, enterovirus D-68, and Gram-negative bacteria that produce carbapenemases. Every effort was made to include up-to-date information in the Manual on these recently emergent organisms. In addition, the studies of the human microbiome have informed our understanding of normal microbial communities and have posed the possibility of polymicrobial rather than single-agent infections. In conclusion, we are profoundly grateful for the privilege of guiding the Manual through the publication of this 11th edition. We hope that the efforts of the editors and authors will prove useful to the clinical microbiology community until the next edition is available in about 4 more years.

JAMES H. JORGENSEN MICHAEL A. PFALLER

xxv

MCM11_FM.indd 25

3/19/15 1:07 PM

Author and Editor Conflicts of Interest

April N. Abbott (coauthor on chapter 77) has participated in research studies supported by Cepheid, ELITech, and Nanosphere.

Ferric C. Fang (coauthor on chapter 77) has participated in research studies supported by Cepheid, ELITech, and Nanosphere.

Ellen Jo Baron (chapters 18, 50) is an employee of Cepheid.

Charlotte Gaydos (coauthor on chapter 63) has received research funding from Hologic/GenProbe, Abbott, Bectin Dickinson, Roche, and Cepheid.

Karen Bush (coauthor on chapter 68) has received retirement compensation from Bristol-Myers Squibb, Johnson & Johnson, and Pfizer (Wyeth). She has served as advisory board member or consultant for or has received research support from Allegra, Apple Tree Partners, AstraZeneca, Basilea, Cempra, Cubist, Fedora, Medivir, Merck, Novartis, Rempex, Roche, Shionogi, The Medicines Company, Theravance, and Wockhardt.

Christine C. Ginocchio (section editor; coauthor on chapters 80, 81, and 104) is an employee of bioMérieux. She has received clinical trial/research funding from Diagnostic Hybrids/ Quidel, Inc. and research funding and Advisory Board fees from Hologic.

Karen C. Carroll (volume editor; coauthor on chapters 4 and 53) has received research funding or has been a consultant for Abbott Molecular, Inc., Accelerate, Inc., AdvanDx, Inc., BD Diagnostics, Inc., BioFire, Inc., Quidel, Inc., Nanosphere, Inc., and NanoMR. She also receives royalties from McGraw-Hill Publishers.

Patti E. Gravitt (coauthor on chapter 104) has received research funding from Roche and Merck and served on an Advisory Board for Qiagen.

Maiken Cavling-Arendrup (coauthor on chapter 131) has received research funding and/or been a consultant and/or invited speaker for Astellas, Cephalon, Gilead Sciences, Merck Sharpe & Dohme, Pfizer, Schering-Plough, and Swedish Orphan.

Randall Hayden (coauthor on chapter 103) is a consultant for Quidel Corporation and has research agreements and collaborations with Focus Diagnostics, Luminex Molecular Diagnostics, Inc., EliTech, and BioFire Diagnostics, Inc.

Sharon Chen (coauthor on chapter 119) has served on antifungal advisory boards of Merck, Gilead Sciences, and Pfizer and received untied grants from Merck, Pfizer, and Gilead Sciences.

Janet A. Hindler (coauthor on chapter 74) has received funding for research from bioMérieux, Siemens Healthcare, and BD Biosciences.

Christopher Doern (coauthor on chapter 5) has served as a consultant to ThermoFisher and Copan and has received research funding from Becton Dickinson, BioFire, and bioMérieux.

Romney M. Humphries (coauthor on chapter 74) has received funding for research from bioMérieux, Siemens Healthcare, and BD Biosciences.

J. Stephen Dumler (section editor; chapter 61; coauthor on chapters 48 and 65) holds a patent for the in vitro cultivation of Anaplasma phagocytophilum that is used as a primary method to prepare serodiagnostic tests. The technology is licensed to Focus Laboratories and he receives periodic royalties for this license.

Elizabeth M. Johnson (section editor; coauthor on chapter 131) has received research funding and/or been a consultant and/or invited speaker for Astellas, Gilead Sciences, Merck Sharp & Dohme, Pfizer, and Schering-Plough.

Patricia Harris (coauthor on chapter 80) is a former employee of Diagnostic Hybrids/Quidel, Inc.

xxvi

MCM11_FM.indd 26

3/19/15 1:07 PM

AUTHOR AND EDITOR CONFLICTS OF INTEREST n  xxvii James H. Jorgensen (co-editor in chief; coauthor on chapters 1 and 71) serves on an advisory board for Accelerate Diagnostics and has received research support from Merck Pharmaceuticals. Daryl M. Lamson (coauthor on chapter 88) has a patent application pending for the discovery of HRV-C sequences and their uses (application number PCT/US 2009/0275636). James S. Lewis, II (coauthor on chapter 68), has served as consultant to Acceler8 Technology, Astellas, Forest, Merck, Pfizer, Trius, Cubist, Rempex, and Theravance and is on the speakers bureaus of Astellas, Forest, and Merck.

Michael A. Pfaller (co-editor in chief; coauthor on chapter 1) is an employee of T2 Biosystems. Bobbi S. Pritt (section editor; coauthor on chapter 136) has received a grant to perform research in collaboration with the University of Minnesota to improve the performance of rapid diagnostic tests, funded through the Minnesota Partnership for Biotechnology and Medical Genomics program. She has also participated in research studies supported by Roche Diagnostics and Gen-Probe.

Stephen E. Lindstrom (coauthor on chapter 84) has a patent on realtime RT-PCR assays for influenza.

Sandra Richter (volume editor; coauthor on chapters 72 and 79) has participated in research studies supported by Achaogen, BD Diagnostics, bioMérieux, BioFire, Forest Laboratories, Nanosphere, Pocared, and OpGen.

Melissa Miller (section editor) has served as consultant to Becton Dickinson, Meridian Biosciences, and GenMark Dx and has received clinical trial and research funding from Becton Dickinson, Luminex Molecular Diagnostics, Cepheid, Hologic, and Nanosphere.

Robert Shafer (coauthor on chapter 111) has consulted for Celera. He has received unrestricted research gifts from Celera, Siemens Health Care, and Hoffman LaRoche and research funding from Merck Pharmaceuticals, Bristol-Myers Squibb, Hoffmann-LaRoche, and Gilead Sciences.

Susan Novak-Weekley (coauthor on chapter 140) was a member of the Roche Advisory Board in 2012.

Jennifer S. Smith (coauthor on chapter 104) has received research funding or advisory board consultation fees from BD, GlaxoSmithKlein, Hologic, Merck, Qiagen, and Trovogene.

Robin Patel (section editor; coauthor on chapter 4) has received research support from Pfizer, Pradama, Astellas, Tornier, bioMérieux, NanoMR, 3M, BioFire, and Bruker. She has patents on an antibiofilm substance, a method and apparatus for device sonication, and a PCR assay for Bordetella pertussis and Bordetella parapertussis. David Perlin (chapter 130) is listed on a pending patent application concerning diagnostics for echinocandin resistance.

MCM11_FM.indd 27

Tania Sorrell (coauthor on chapter 119) has served on antifungal advisory boards of Merck, Gilead Siences, and Pfizer and received untied grants from Merck, Pfizer, and Gilead Sciences. Yi-Wei Tang (section editor; coauthor on chapters 81 and 86) has research agreements and collaborations with Alere, Cepheid, Luminex, and Roche.

3/19/15 1:07 PM

MCM11_FM.indd 28

3/19/15 1:07 PM

Diagnostic Strategies and General Topics VOLUME EDITOR: SANDRA S. RICHTER SECTION EDITOR: ROBIN PATEL

1

2

Introduction to the 11th Edition of the Manual of Clinical Microbiology / 3

9

JAMES H. JORGENSEN AND MICHAEL A. PFALLER

10 Molecular Epidemiology / 131

Microscopy / 5 DANNY L. WIEDBRAUK

3

Laboratory Detection of Bacteremia and Fungemia / 15 MICHAEL L. WILSON, MELVIN P. WEINSTEIN, AND L. BARTH RELLER

4

Systems for Identification of Bacteria and Fungi / 29 KAREN C. CARROLL AND ROBIN PATEL

5

6

Automation and Design of the Clinical Microbiology Laboratory / 44

EIJA TREES, PAUL A. ROTA, DUNCAN MACCANNELL, AND PETER GERNER-SMIDT

11 Procedures for the Storage of Microorganisms / 161 ROSEMARY C. SHE AND CATHY A. PETTI

12 Prevention of Laboratory-Acquired Infections / 169 MICHAEL A. NOBLE

13 Decontamination, Disinfection, and Sterilization / 183 CONSTANZE WENDT, RENO FREI, AND ANDREAS F. WIDMER

14 Biothreat Agents / 217

Molecular Microbiology / 54

15 The Human Microbiome / 226

Immunoassays for Diagnosis of Infectious Diseases / 91 ELITZA S. THEEL, A. BETTS CARPENTER, AND MATTHEW J. BINNICKER

8

IONA MUNJAL AND BELINDA OSTROWSKY

CHRISTOPHER D. DOERN AND MARTIN HOLFELDER FREDERICK S. NOLTE

7

Investigation of Disease Outbreaks / 120

Prevention of Health Care-Associated Infections / 106 DANIEL J. DIEKEMA AND MICHAEL A. PFALLER

SUSAN E. SHARP AND MICHAEL LOEFFELHOLZ JAMES VERSALOVIC, SARAH K. HIGHLANDER, AND JOSEPH F. PETROSINO

16 Microbial Genomics and Pathogen Discovery / 238 JENNIFER K. SPINLER, PEERA HEMARAJATA, AND JAMES VERSALOVIC

This page intentionally left blank.

Introduction to the 11th Edition of the Manual of Clinical Microbiology* JAMES H. JORGENSEN AND MICHAEL A. PFALLER

1 The 11th edition of the Manual of Clinical Microbiology (MCM11) marks a significant change from prior editions in that there are co-editors in chief for the first time. There has been only one editor in chief for all prior editions since the first edition in 1970. However, as the Manual has grown in size and scope, it has become a Herculean effort for a single editor in chief. Since the 8th edition of the Manual, it has been published as two volumes. We have divided our duties primarily by volume: volume 1 (Jorgensen) and volume 2 (Pfaller). This edition of the Manual contains 151 chapters comprising almost 2,600 pages. It includes a comprehensive array of content on all aspects of microbiology contributed by world experts on each subject, with their chapters edited by a highly committed team of volume and section editors working within the usual 4-year publication cycle. This edition includes one new volume editor (Sandra Rich-ter) and three new section editors in addition to the new editors in chief. Four volume editors, Karen Carroll, Guido Funke, Marie Louise Landry, and David Warnock, served for the past edition of the Manual. Underscoring the international importance of the Manual, 20% and 37% of section editors and chapter authors, respectively, contributed content from countries outside the United States (up from 19 and 30% in the previous edition). There were 88 authors who contributed content to the Manual for the first time. The overall organization of the Manual and the chapter formats are quite similar to those of the 10th edition. However, readers will note some new or expanded content in this edition. Since publication of the 10th edition, the application of matrix-assisted laser desorption ionization– time of flight mass spectrometry (MALDI-TOF MS) has rapidly been developed and embraced by clinical microbiology laboratories for identification of a vast array of bacteria and fungi. Most of the “organism” chapters now include comments on the utility of MALDI-TOF MS for the particular genera and species under discussion. Microbiologists have quickly joined this bandwagon due to the speed, accuracy of identifications, and low cost per test of this technique (despite the high initial instrument acquisition cost). The technology is still evolving, as the required extensive databases needed for all of the relevant organism groups are being painstakingly constructed. Assuming the adequacy of

the databases, this technology may supplant much of the conventional phenotypic testing of the past and even the use of sequence-based identification approaches. Having said that, the cost and complexity of gene sequencing has continued to decrease rapidly, making it likely to be accessible to more clinical microbiology laboratories in the next few years. We have learned that no one genomic, proteomic, or phenotypic approach is perfect for the identification of all organisms, and different approaches may need to be used in concert to accurately identify some organisms. Readers will note many new genus and species names in this edition. These result from initial descriptions of some species and reclassification of some genera and species from a previous taxon to a new or different one based upon 16S or 23S gene sequencing studies, sequencing of certain housekeeping genes, or DNA hybridization efforts that have demonstrated new phylogenetic associations. Some of these may seem bewildering at first due to the many new names that are unfamiliar to both microbiologists and practicing clinicians. Some of the newly designated organisms may not be recognized by conventional phenotypic testing and may not be found in the databases of FDA-cleared identification devices. They may require gene sequencing, use of microarrays, or perhaps mass spectrometry for accurate identifications. This will put pressure on clinical microbiology laboratories to either adopt newer technologies or place greater reliance on reference laboratories when it is important to know the contemporary identities of significant organisms. This offers an opportunity for dialogue between clinical microbiologists and the clinicians that they serve regarding the situations in which intensive identification efforts are justified. The structure of volume 2 is largely the same as that found in MCM10. In Section IV (Virology), the chapter on Hendra and Nipah viruses from the previous edition has been folded into chapter 85 (“Parainfluenza and Mumps Viruses”) and also includes other members of the Paramyxoviridae, and chapter 106 now highlights bocaviruses along with parvoviruses to recognize the emergence of the former as a human pathogen since its discovery in 2005. The Mycology and Parasitology sections remain largely unchanged, with the exception of the movement of the chapter on microsporidia from the Parasitology section to the Mycology section in recognition of the reclassification of these agents to the kingdom Fungi. As with the Bacteriology chapters,

*This chapter contains some information presented in chapter 1 by James Versalovic in the 10th edition of the Manual.

doi:10.1128/9781555817381.ch1

3

4

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

the chapters in volume 2 reflect the tremendous advances in the area of molecular taxonomy which have resulted in the reclassification of many different organisms, the recognition of several so-called “cryptic” species, and the discovery of several “new” pathogens as molecular and proteomic tools find their way from the research setting to the clinical laboratory. Now entering its 6th decade, the Manual strives to continue to be the leading, most authoritative reference for the “real-world” practice of clinical microbiology. In order to create and assemble each edition, this publication builds on the content of past editions, and the process requires about 3 years of careful planning, design, writing, and review of chapters before the final phases of copyediting, composition, printing, and binding. In the intervening 1 to 2 years from the time of chapter acceptances until printing, new

diagnostic trends, technologies, pathogens, and patterns of infectious diseases may emerge or change in ways that affect the timeliness and relevance of this comprehensive reference. This sobering reality simply “goes with the territory” of compiling any authoritative body of work. Hopefully the Manual continues to provide a highly respected benchmark and authoritative reference for the entire field of clinical microbiology. In the era of mass collaboration and rapid communication, our team at the Manual trusts that our readership, each of you, will contribute to the future of this field by pointing out errors, issues, and trends that serve to strengthen the Manual and its next edition. The work never stops, and the knowledge base keeps growing. So let us all continue to enhance the practice and contribute to the evolution of our cherished profession of clinical microbiology.

Microscopy DANNY L. WIEDBRAUK

2 The history of microscopy has been closely linked to the beginning of microbiology since 1665, when Hooke published his treatise Micrographica, which included illustrations of mold forms and the anatomy of the flea (1). Today, light microscopy is used not only in microbiology, pathology, and cell biology but also in metallurgy, materials science, computer chip design, and microsurgical applications. This chapter will attempt to describe the basic concepts of light microscopy as they are practiced in the microbiology laboratory.

that appear to be in sharp focus. Depth of field decreases as the numerical aperture (NA) of the lens increases (4). Depth of focus is the area around the image plane where the image will appear to be sharply focused. The image plane is formed within the microscope tube at or near the level of the ocular lenses. Microscopes with greater depths of focus allow the user to employ ocular lenses with different working distances, magnification factors, and visual compensation systems without losing image sharpness. Like depth of field, depth of focus depends upon the numerical aperture of the objective. However, depth of focus increases as the numerical aperture increases (4).

TECHNICAL BACKGROUND AND DEFINITION OF TERMS

Immersion Fluid (Immersion Oil) Immersion fluid is a term used to describe any liquid that occupies the space between the object and microscope objective lens. Immersion fluids are usually required for objectives that have working distances of 3 mm or less (2). Many microscopy applications employ immersion fluids that possess the same refractive index as the glass slide (refractive index = 1.515) (2, 4). This procedure produces a homogeneous optical path which minimizes light refraction and maximizes the effective numerical aperture of the objective lens. Immersion fluids are also used between the condenser and the microscope slide in transmitted light fluorescence microscopy and in dark-field microscopy to minimize refraction, to increase the numerical aperture of the objective, and to improve optical resolution (2, 4).

Aberration Aberrations are unwanted artifacts in the microscopic image that are caused by elements in the optical path. Aberration can be caused by physical objects, such as dust or oils, on the optical surfaces, by alterations in the light path caused by improper alignment or aperture settings, and by lens system imperfections. Two main types of optical aberration, spherical aberration and chromatic aberration, can occur when white light passes through a convex lens. Spherical aberration is exhibited by images that appear to be in focus in the center of the field and out of focus at the periphery (2). Chromatic aberration occurs because shorter light wavelengths are refracted to a greater extent than longer wavelengths (2). This wavelength separation (also called dispersion) produces color fringes within the image field. Lenses that are not corrected for chromatic aberration can cause difficulties when interpreting Gram and other staining due to the presence of purple or green fringes around bacteria. This is especially problematic when the bacteria are very small.

Köhler Illumination

Contrast is a measure of the differences in image luminance that provide gray scale or color information. Contrast is expressed as the ratio of the difference in luminance between two points divided by the average luminance in the field (3). Under optimum conditions, the human eye can detect the presence of 2% contrast (3).

Köhler illumination was first introduced in 1893 by August Köhler of the Carl Zeiss Corporation as a method for providing the optimum specimen illumination (2). In this procedure, the collector lens projects an enlarged and focused image of the lamp filament onto the plane of the aperture diaphragm. Because the light source is not focused at the specimen, the specimen is bathed in a uniformly bright, glare-free light that is not seriously affected by dust and imperfections on the glass surfaces of the condenser. Köhler illumination is required to produce the maximum optical resolution and high-quality photomicrographs (2, 5).

Depth of Field and Depth of Focus

Mechanical Tube Length

Depth of field is a subjective measure of the vertical distance between the nearest and farthest objects in the specimen

Mechanical tube length describes the light path distance within the microscope body tube. Tube length is measured

Contrast

doi:10.1128/9781555817381.ch2

5

6

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

FIGURE 1 Objective lens labeling. Objective lenses are labeled with information about the manufacturer, correction factors, numerical aperture, tube length, coverslip thickness, working distance, and expected immersion medium. Objectives without a listed aberration correction are considered achromats. Objectives without a listed immersion medium (Oil, Oel, W, Gly) are considered dry objectives and are intended for use with air between the lens and the specimen. doi:10.1128/9781555817381.ch2.f1

from the objective opening in the nosepiece to the top edge of the observation tube. Tube length is usually inscribed on the barrel of the objective as the length in millimeters (160, 170, 210 mm, etc.) for fixed lengths or the infinity symbol (∞) for infinity-corrected tube lengths (Fig. 1). Many of the newer objectives are infinity corrected, while older objectives are corrected for 160-mm (Nikon, Olympus, Zeiss) or 170-mm (Leica) tube lengths (2).

Numerical Aperture NA is a measure of the light-gathering capability of a lens or condenser. Higher NA objectives have better resolving power and brighter images than lower NA objectives. Higher NA objectives also have a shallower depth of field. The equation for determining NA is given by NA = n × sin(θ), where n is the refractive index of the imaging medium between the objective and the specimen and θ is one-half the angular aperture of the objective (Fig. 2) (1, 2, 4).

Refractive Index (Index of Refraction) Index of refraction is the ratio of the velocity of light in a vacuum to its velocity in a transparent or translucent medium (2, 4, 6). As the refractive index of a material increases, light beams entering or leaving a material are deflected (refracted) to a greater extent. The refractive index of a medium depends upon the wavelength of light passing through it. Light beams containing multiple wavelengths (e.g., white light) are dispersed when they move into a different medium because the wavelengths in the beam are refracted to slightly different degrees. Light dispersion causes chromatic aberration in microscope objectives (2). Refractive index is also an important variable in calculating numerical aperture (see “Numerical Aperture” above). Moving from a high dry microscope objective that uses air as the imaging medium (refractive index of air = 1.003) to an oil immersion objective of the same power (refractive index of immersion oil = 1.515) increases the maximum theoretical numerical aperture of a given lens from 1.0 to 1.5, producing a 50% increase in light-gathering capability (2, 4).

FIGURE 2 Typical configuration for bright-field microscopy. The column of light generated by the field lens and the field diaphragm enters the bottom of the condenser and is focused on the slide by the condenser lens. The condenser diaphragm controls the angle of the light, the numerical aperture of the condenser, and the amount of contrast in the image. The working distance is the vertical distance from the top of the specimen to the leading edge of the objective lens. The semiangle of the objective aperture (θ) is used to calculate numerical aperture. Modified from reference 4. doi:10.1128/9781555817381.ch2.f2

Resolution (Resolving Power) The resolution of an optical microscope is defined as the shortest distance between two points that can be distinguished as separate entities by the observer or camera system (4). The resolving power of a microscope is the most important feature of the optical system because it defines our ability to distinguish fine details in a specimen. The theoretical limit of resolution (r) for a given lens is defined mathematically as r = κ/(2NA), where κ is the imaging wavelength and NA is the numerical aperture of the lens (4). From this equation, it is obvious that only the light wavelength and NA directly affect the resolving power. Thus, a 40× oil objective with an NA of 1.30 can have the same resolving power as a 100× oil objective (Table 1). In the same manner, the resolving power of a 100× oil objective will be higher when ultraviolet (UV) wavelengths are used than when visible light is used (Table 1).

Working Distance Working distance is the distance between the leading edge of the objective lens and the top of the cover glass when the specimen is in focus (Fig. 2). The working distance of an objective generally decreases as magnification increases (4). The working distance of an objective may not be inscribed on the barrels of older objectives, but newer objectives often contain the working distance in millimeters (Fig. 1). Longer working distance objectives are important when examining the inside surfaces of glass tubes (tube cultures) and cell culture flasks.

SIMPLE MICROSCOPE Common objects, such as jeweler’s loupes, photographic slide viewers, and simple magnifying or reading glasses, are all examples of simple microscopes. A simple microscope contains a single bi-convex magnifying lens which is

2. Microscopy n TABLE 1

7

Resolving the power of selected lenses with different numerical apertures.

Lens system Eye Hand magnifier 10× objective 40× objective 40× objective (oil) 100× objective 100× objective

NA

Light color Avg wavelength (nm)

Medium

0.03 0.30 0.75 1.30 1.30 1.30

White White White White White White UV

Air Air Air Air Oil Oil Oil

550 550 550 550 550 550 400

Resolution (μm) 700 10 0.92 0.37 0.21 0.21 0.15

thicker in the center than at the periphery. In contrast, with compound microscopes, simple microscopes produce a magnified image that is in the same orientation as the original object. Because of their low NA, simple microscopes have limited resolution and magnifying power. Most commercial magnifiers are able to produce a ×2 to 30 magnification, and the better lenses will have a resolution of about 10 μm. Simple magnifiers are useful for dissection, examination of bacterial colonies, and interpretation of agglutination reactions.

provide a white light. Newer microscopy systems now use light-emitting diode (LED) light matrices. These systems have reduced energy requirements, do not heat the specimen or the lower optical train, and last for decades. Light generated by the light source is passed through a collector and a field lens (Fig. 3) before being directed into the substage condenser and onto the specimen. Imageforming light rays are captured by the microscope objective and passed into the eyepieces or a camera port. Alignment of the optical components of a microscope is critical to producing a good image.

COMPOUND MICROSCOPE

Field Diaphragm

The first compound microscopes were constructed around 1590 by Dutch spectacle makers Zaccharias Janssen and Hans Janssen. The Janssen microscope consisted of an object lens (objective) that was placed close to the specimen and the eye, or an ocular lens that was placed close to the eye. The lenses were separated by a body tube. In this microscope, the objective lens projected a magnified image into the body tube and the eyepiece magnified the projected image, thereby producing a two-stage magnification. Modern compound microscopes still use this general design and have two separate lens systems mounted at opposite ends of a body tube. The stereoscopic microscope combines two compound microscopes, which produce separate images for each eye. The three-dimensional stereoscopic effect is produced in the brain when two images are viewed with slightly offset viewing perspectives. Placing a camera on one of the eyepieces or on a dedicated camera port will not produce a three-dimensional photo image. Stereoscopic microscopes are used with reflected or transmitted illumination, but the absence of a substage condenser limits their NA and resolution. Stereomicroscopes are useful in examining the colonial morphology of bacteria, fungi, and cell cultures.

The field diaphragm is located in the light path between the light source and the substage condenser (Fig. 3). This iris-like mechanism controls the width of the light beam that enters the substage condenser. The field diaphragm does not affect the optical resolution, numerical aperture, or intensity of illumination. However, the field diaphragm should be centered in the optical path and opened far enough that it just overfills the field of view. This adjustment is important for preventing glare and loss of contrast in the observed image. When the field diaphragm is opened

Optical Train The modern light microscope is composed of optical and mechanical components that, together with the mounted specimen, make up the optical train. The optical train of a typical bright-field microscope consists of an illuminator (light source and collector lens), a substage condenser, a specimen, an objective, the eyepiece, and a detector. The detector can be a camera or the observer’s eye. Specimen illumination is one of the most critical elements in optical microscopy. Inadequate or improper sample illumination can reduce contrast in the specimen and significantly decrease the resolving power of any microscope (7). Fifty or 100-watt tungsten halogen lamp systems have been the most popular means of providing light for visible-light microscopy because they have a relatively low cost and

FIGURE 3 Anatomy of a typical clinical microscope with an integral camera. doi:10.1128/9781555817381.ch2.f3

8 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

too far, scattered light and reflections can degrade image quality (1).

Substage Condenser The substage condenser is typically mounted beneath the microscope stage in a bracket that can be raised or lowered independently of the stage (Fig. 3). The substage condenser gathers light from the field diaphragm and concentrates it into a cone of light that illuminates the specimen with uniform intensity over the entire field of view. Adjustment of the substage condenser is probably the most critical element for achieving proper illumination, and it is the main source of image degradation and poor-quality photomicrography. The condenser light cone must be properly adjusted to optimize the intensity and angle of light entering the objective. Because each objective has different light-gathering capabilities (numerical aperture), the substage condenser should be adjusted to provide a light cone that matches the numerical aperture of the new objective. This is done by adjusting the aperture (or condenser) diaphragm control. Substage condensers on newer microscopes have a scale embossed on the condenser and an index mark on the aperture control that allows the user to quickly switch from one NA range to another. Many manufacturers are now synchronizing the NA gradations to correspond with the approximate numerical aperture of the objectives. In clinical laboratory practice, the condenser aperture is often made smaller to improve the contrast of wet mounts and some stained preparations (1). This practice, while effective for some applications, will result in decreased resolution (3). It should be noted that the intensity of illumination should not be adjusted by opening and closing the condenser aperture diaphragm or by moving the condenser laterally in the light path. Illumination intensity should be controlled through the use of neutral density filters placed into the light path or by reducing voltage to the lamp. It should be noted that reducing the voltage will also alter the color of the light emanating from a tungsten halogen bulb (5). The lighting intensity provided by LED light systems can be reduced without altering the color of the light. The color of the incoming light will affect photomicroscopic color balance, and it can influence the interpretation of stained specimens.

Objectives The objective lens is the most important single determinant of the quality of the image produced by a particular microscope (1). When choosing a microscope, the purchaser must select the magnification factor, the NA, and the level of correction for each objective. Lenses with higher NA values will have higher resolution and produce a brighter field of view. Choosing an appropriate level of optical correction will depend upon the ultimate use of the microscope. Achromatic (achromat) objectives are the least expensive objectives found on laboratory microscopes. Achromat objectives are corrected for axial chromatic aberration in two wavelengths (red and blue), and they are corrected for spherical aberration in one color (green) (2). The limited correction of achromatic objectives can cause a number of optical artifacts when specimens are examined and photographed in color (e.g., green images often have a reddish-magenta halo) (2). Achromat objectives produce the best results when the light passes through a green filter and when blackand-white photomicroscopy is performed. Flatness of field

is also a problem when using straight achromat objectives because the center of the field is in focus while the edges are out of focus (2). In the past few years, most manufacturers have begun providing flat-field corrections for achromat objectives. These objectives are called plan-achromats. The next-higher level of correction and cost is found in objectives called fluorites or semiapochromats. Fluorite objectives are produced from advanced glass formulations that allow for greatly improved correction of optical aberration. Like achromat objectives, fluorite objectives are corrected chromatically for red and blue light (2). Fluorites, like achromats, are corrected spherically for two or three colors instead of a single color (2). The superior correction of fluorite objectives compared to that of achromat objectives enables these lenses to be made with a higher numerical aperture. Fluorite lenses produce brighter images than achromats. Fluorite objectives also have better resolving power than achromats and provide a higher degree of contrast, making them better suited for color photomicrography in white light (2, 4). Apochromats are the most highly corrected microscope lenses and the most costly. Apochromats are corrected chromatically for three colors (red, green, and blue), which almost eliminates chromatic aberration, and are corrected spherically for either two or three wavelengths (2). Apochromat objectives are the best choice for color photomicrography in white light. Because of their high level of correction, apochromat objectives usually have, for a given magnification, higher numerical apertures than do achromats or fluorites (2, 4). Fluorescence objectives are designed with quartz and other special glasses that have high rates of transmission of UV, visible, and infrared light. These objectives are extremely low in auto-fluorescence and use specialized optical cements and antireflection coatings that protect the lens and allow it to operate with a wide variety of excitation wavelengths. Correction for optical aberration and numerical aperture values in UV fluor objectives usually approaches that of apochromats, which contributes to image brightness and enhanced image resolution (2, 8). The primary drawback of high-performance fluorescence objectives is that many are not corrected for field curvature and produce images that do not have uniform focus throughout the entire field of view. This is not a large problem when performing direct or indirect fluorescent antibody testing but it can be troublesome if you have to use the same objectives for bright-field or phase-contrast microscopy. Microscope objectives that use air as the medium between the coverslip and the objective lens are considered dry objectives. The maximum working numerical aperture of a dry objective system is limited to 0.95, and greater values can be achieved only with optics designed for immersion media. Immersion media have the same refractive index and dispersion values as glass (refractive index = 1.51). The use of immersion media produces a homogeneous light path from the coverslip to the lens so that light is not refracted away from the objective. The use of immersion fluids and lenses significantly increases the numerical aperture and the optical resolution of the system. In addition to oil lenses, specially corrected objective lenses designed for glycerin and water immersion are available commercially. The proper immersion fluid type is always stamped on the side of the objective. The advantages of oil immersion objectives are severely compromised if the wrong immersion fluid is utilized. Microscope manufacturers produce immersion objectives with tight refractive index and dispersion tolerances (2). It is therefore advisable to use only the immersion

2. Microscopy n 9

fluid recommended by the objective manufacturer. Mixing of immersion fluids from different manufacturers should be avoided because mixing can produce unexpected crystallization artifacts or phase separations that compromise image quality. Many high-power (NA ≥ 0.8) dry objectives are engineered to operate through 0.17-mm coverslips (designated number 1 1/2). In practice, however, the total thickness of the specimen/coverslip sandwich can be greater or less than 0.17 mm due to variations in coverslip and/or mounting fluid thickness (2, 4). Under these conditions, there will be noticeable spherical aberration in the microscopic image (2, 4). A 0.2-mm deviation in coverslip thickness will produce an 8% decrease in image intensity with a 0.79-NA objective and a 57% decrease with a 0.85-NA high dry objective (2). Therefore, some of the more advanced dry objectives are engineered with a coverslip correction collar that adjusts the objective lens elements to compensate for coverslip thickness variations. Objectives with a coverslip correction collar are labeled “Corr,” “w/Corr,” or “CR.” However, this labeling is usually unnecessary because the objective has a distinctive knurled ring and graduated scale on the side. The expected coverslip thickness for an objective is etched on the barrel of the objective (Fig. 1). The eyepiece or ocular objective contains the final lens system in the optical train. The purpose of the ocular objective is to magnify and focus the projected image onto the eye of the viewer. Ocular lenses generally have a magnification factor of ×10 to ×20, and the total magnification of the microscope is the product of the objective magnification and the ocular magnification (1, 2, 6). Thus, a microscope with a 40× objective and a 10× ocular would have a magnification value of ×400. Many eyepieces have a shelf at the level of the fixed eyepiece diaphragm that allows for the insertion of ocular micrometers, pointers, or crosshairs. This shelf is located at the focal plane of the image projected by the objective lens so that the inserted element is in focus when the specimen image is in focus. When upgrading microscope systems, the normal tendency is to save old objectives and oculars as backups. Check with your microscope manufacturer before doing this. Oculars, especially those from a different manufacturer, can have a different focal length than the one used in the new microscope. In addition, many manufacturers include optical corrections in the oculars that compensate for aberrations in their objective line. These corrections may cause unwanted chromatic aberrations when the old lens is used with the new microscope. Objectives with a standard tube length cannot be used on infinity-corrected optical systems.

DARK-FIELD MICROSCOPY Dark-field microscopy is a specialized illumination technique used to detect thin organisms, such as spirochetes and Leptospira spp. High-resolution dark-field microscopy utilizes a specialized high-NA cardioid dark-field condenser that blocks the central light path light and produces a hollow cone of illumination that is directed away from the objective lens at an oblique angle (Fig. 4). Bacteria on the slide have a slightly different refractive index than the surrounding medium, and light rays passing through the organism are refracted into the objective lens, producing bright organism profiles against a dark background. Darkfield microscopy requires careful alignment of the condenser and placement of immersion oil between the slide and the substage condenser. Dark-field microscopy, when done correctly, increases the resolution of the microscope to 0.1 μm

FIGURE 4 Dark-field illumination. The central light path interacts with the silvered dome located at the bottom of the condenser and is reflected away from the specimen. Peripheral light is reflected into the condenser, and it is reflected again by the internal condenser surfaces to produce a cone of light that is directed obliquely away from the objective. doi:10.1128/9781555817381.ch2.f4

or less (3). The resolution of bright-field microscopy is 0.2 μm (1).

PHASE-CONTRAST MICROSCOPY Many unstained biological specimens are virtually transparent when observed under bright-field illumination. To improve visibility in wet mounts and cell cultures, microscopists often reduce the opening size of the substage condenser iris diaphragm, but this maneuver is accompanied by a serious loss of resolution and the introduction of diffraction artifacts (2, 3). Phase-contrast microscopy significantly improves the contrast in these specimens without significant loss in resolution (3). In phase-contrast microscopy, a ring annulus is placed directly under the lower lens of the condenser to produce a hollow cylinder of light. This light is essentially unchanged as it passes into the objective, and it arrives at the rear focal plane of the objective in the shape of a ring. Light that goes through the specimen is refracted and slowed slightly so that it is out of phase with the unchanged light by about 1/4 wavelength. This light is spread over the entire focal plane. Light passing through the rear focal plane of the objective interacts with a ring-shaped phase plate that alters the direct light path by 1/4 wavelength (3). When the direct light and the refracted light arrive at the image plane, they are out of phase by 1/2 wavelength. This out-of-phase light interacts destructively, so that specimen details appear as dark areas against a lighter background (3). Because the phase-shifting calculations are based upon on a 1/4 wavelength of green light, the phase image has the best resolution when a green filter is placed in the light path (3). Green filters also allow the microscopist to use less expensive achromat lenses that are spherically corrected for green light. Phase microscopy is an important tool for examining living and/or unstained material in wet mounts and cell cultures. However, phase-contrast microscopy has lower resolution than bright-field microscopy of stained specimens (3). In addition, viewed objects are often surrounded by halos that can obscure boundary details. Phase-contrast microscopy does not work well with thick specimens because the phase shift may be greater than the expected 1/4 wavelength.

10

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

FLUORESCENCE MICROSCOPY The fluorescence microscope was developed in the early 1900s, and many of the initial microscopic studies involved identification and localization of compounds that autofluoresced when irradiated with UV light. In the 1930s, a number of investigators began using fluorescent compounds to identify specific tissue components and infectious agents that did not autofluoresce (8). Examples of this type of stain include acridine orange (intercalates into DNA and RNA), auramine-rhodamine (for mycolic acids), Calcofluor White (for fungal cell wall polysaccharides), Evans blue (for the cytoplasm of fixed cells), and Hoechst 33258 (for the minor groove of AT-rich double-stranded DNA). The use of fluorochrome-antibody conjugates (immunofluorescence) was first described in the 1940s, when Coons et al. (9, 10) used fluorescein-labeled antibodies to detect pneumococcal polysaccharide antigens in tissue sections of infected mice. Fluorescent-antibody staining expanded significantly with the development of fluorescein isocyanate in 1950 (11) and the more stable fluorescein isothiocyanate (FITC) derivative in 1958 (12–16). Quantum dots have recently emerged as a new class of fluorescent labels for biology and medicine (17). When conjugated to antibodies and other biological ligands, these tiny light-emitting particles can overcome many of the disadvantages of traditional fluorophores used in clinical pathology. Quantum dots have superior signal brightness, they are resistant to photobleaching, and multiple fluorescent colors can be excited by the same wavelengths of light. The last property makes multicolor fluorescence microscopy easy to perform and, when the wavelengths are tuned properly, provides quantitative information about ligand abundance (18). Quantum dots have been used for staining live cells (18), fixed cells (17), and tissues (17). Today, fluorescence microscopy is also used in conjunction with nucleic acid hybridization to visualize the locations of fluorescent in situ hybridization (FISH) and multicolor FISH probes (19, 20). Fluorescence microscopy is dependent upon the ability of fluorescent substances to absorb near-UV energy and reemit that energy (light) at a lower wavelength (8). To work properly, the fluorescence microscope must irradiate the specimen with UV excitation light and separate the much weaker emitted light from the brighter excitation light so that only the emitted light reaches the eye. The resulting image consists of brightly shining areas against a dark background (8). Older fluorescence microscopes are configured for dark-field illumination or transmitted light fluorescence (8). These systems were cumbersome to use and lacked resolution. Most modern fluorescence microscopes use reflected light (epifluorescence). In these instruments, the excitation light is directed downward through the objective and onto the specimen. The emitted light and the reflected excitation light are collected by the objective, and they pass through a dichromatic mirror, which removes the excitation light and allows the longerwavelength emitted light to form an image. With epifluorescence, the objective acts as a condenser and the alignment and oiling issues associated with a dark-field condenser are eliminated (8). The visual field is brighter with epifluorescence, the resolution is higher, and fluorescence quenching occurs only in the field of view (8). Fluorescence microscopy requires high levels of illumination because the quantum yield of most traditional fluorochromes is low. The most common lamps used in fluorescence microscopy are mercury vapor (HBO) lamps

TABLE 2 Excitation and emission wavelengths of commonly used fluorochromesa Fluorescent compound Acridine orange (singlestranded nucleic acid) Acridine orange (doublestranded nucleic acid) Auramine O Calcofluor White Ethidium bromide Evans blue Fluorescein isothiocyanate (FITC) Hoechst 33258 Rhodamine B Tetramethylrhodamine isothiocyanate (TRITC)

Excitation wavelength (nm)

Emission wavelength (nm)

500

526

460

640

460 440 545 550

550 500–520 605 610

490

525

352 540

461 625

555

580

a Excitation and emission wavelengths can vary depending upon the solvent and the pH of the solution.

ranging in wattage from 50 to 200 watts or Xenon vapor (XBO) lamps that range from 75 to 200 watts. It should be noted that lamp wattage is not necessarily a measure of usable brightness in a fluorescence lamp. The 100watt HBO lamp is 4 times brighter than the 200-watt HBO lamp and 11 times brighter than the 150-watt XBO lamp (8). HBO and XBO lamps are under high pressure, and care must be taken to prevent the lamps from exploding. One should never touch these lamps with bare hands because oils on the fingers can etch or discolor the glass. Fluorochromes must be excited by specific light wavelengths in order to generate the maximum amount of emitted light. Therefore, specific exciter and barrier filter combinations are used to maximize the quantum yield of the fluorophore. Exciter filters are used to select the required light wavelengths from the spectrum of light generated by the lamp (8). Excitation filters are provided in narrow, medium, and wide bandpass configurations that pass narrow, midsize, and wide ranges of light frequencies, respectively. Barrier filters block shorter light wavelengths and allow longer wavelengths to pass through the filter. Barrier filters are important because they remove the high-intensity excitation light, which can overwhelm the low-intensity emitted light. Barrier filters also prevent UV light from entering the eye, where it can cause cataracts and retinal damage. Wide-bandpass barrier filters generally produce brighter images, but care must be taken to prevent the introduction of background light, which can overwhelm the emitted light. Epifluorescence microscopes also have a dichromatic mirror (beam splitter) that reflects the incoming excitation light to the objective and allows the emitted light to pass to the barrier filter and on to the objectives (7). In most modern epifluorescence microscopes, the barrier filter, excitation mirror, and beam splitter are housed in removable optical blocks, and several of these blocks can be installed in the microscope at one time. This configuration allows the user to quickly change the excitation and barrier filters to accommodate different fluorochromes. Care must be exercised when selecting optical blocks. The excitation filter should match the excitation wavelength of the fluorophore (Table 2), and the emission barrier should allow the emitted light to

2. Microscopy n

pass through. For example, direct fluorescent-antibody testing for viral antigens in cell smears typically employs FITC-labeled antibodies and an Evans blue counterstain. Choosing an optical block with a 450- to 490-nm excitation filter and a 515-nm wide-bandpass barrier filter will produce a bright field of view, and the counterstained cells will appear orange-red. By selecting a more restricted bandpass barrier filter (520 to 560 nm), the field of view will be darker and the red emitted light from the Evans blue counterstain will not be visible. The images produced by this optical block will have more contrast because the background is darker. Both filter combinations are appropriate for this task, but the final choice will depend upon user preference. One of the major problems in the use and examination of fluorescent microscopic images is the tendency of traditional fluorophores to lose fluorescence when exposed to excitation light for several minutes. This loss of fluorescence is caused by two mechanisms, photobleaching and quenching. Photobleaching (fading) is a permanent loss of fluorescence that is caused by chemical damage to the fluorophore (8). Quenching is caused by the presence of free radicals, salts of heavy metals, or halogen compounds (8). Quenching can also be caused by transfer of emission light energy to other fluorescent molecules in close proximity to the fluorophore in a process called fluorescent resonance energy transfer (FRET). To lessen the effect of quenching, slides should be stored in the dark at 2 to 8°C. In addition, the user should block the excitation light path when not viewing or photographing the specimen. Most epifluorescence microscopes have a shutter in the light path for this purpose. Quenching can be a significant problem when photographing fluorescent images because the shutter may be open for a minute or more. Quenching can be reduced somewhat by adding free-radical scavengers, such as p-phenylenediamine (14), 1,4-diazabicyclo(2,2,2)-octane (DABCO) (21), or n-propyl gallate (22) to the mounting fluid. p-Phenylenediamine and n-propyl gallate can be used to reduce quenching in FITC and rhodamine. DABCO is slightly less effective than p-phenylenediamine for FITC fluorescence, but unlike p-phenylenediamine, DABCO does not darken when exposed to light and it is safer to use. Quench-resistant mounting media are also available from Vector Laboratories (Burlingame, CA), Molecular Probes Inc. (Carlsbad, CA), and Bio-Rad Laboratories (Hercules, CA). The major advantage of quantum dot fluorophores is their resistance to photobleaching.

LINEAR MEASUREMENTS (MICROMETRY) The first reported micrometric procedures were credited to Antonie van Leeuwenhoek, who used fine grains of sand as a gauge to determine the sizes of human erythrocytes. Since then, a variety of methods have been used to determine the dimensions of microscopic organisms. The crudest method involves comparing the object size to the measured or calculated view field size. Another rough micrometric method is to compare the sizes of larger organisms to the size of a red blood cell (6 to 8 μm) in the image. Other micrometric techniques include the addition of polystyrene beads of known size into the specimen. Comparative measurements are then performed by utilizing a photomicrograph or digital image. The accuracy of these methods is variable and depends on the homogeneity of the comparison objects.

11

Microorganisms can be measured directly by placing them on a calibrated microscope slide or a counting chamber. The accuracy of this method depends upon the separation distance between ruled lines, but it averages between 10 and 50 μm. The most common procedure used in the clinical laboratory utilizes a graduated scale (reticle) located within one of the eyepieces (23). Reticles must be calibrated against a stage micrometer for each objective (23). To avoid unnecessary recalibrations, the calibration information for each objective should be recorded and stored near the microscope workstation. The accuracy of reticle measurement is approximately 2 to 10 μm (3 to 5%), depending on magnification and the resolution of the stage micrometer (23).

PHOTOMICROSCOPY Microscopists began capturing microscopic images on film shortly after the photographic process was invented (5). Micrographic images have long been used for investigations of morphology, in scientific publications and lectures, and in teaching. Modern film technologies have high resolution and clarity, but the use of photomicrographs in day-today microscopy has been hampered by long turnaround times associated with film development and printing. Reacquiring fluorescence images is a particular concern because the fluorescence can fade (8). The availability of high-quality digital cameras has significantly changed how photomicrographs are used in the microbiology laboratory. Today it is not unusual for digital photomicrographs to be shared with experts via the Internet. This process significantly extends the capabilities of the on-site microbiologist and can enhance patient care. Microscope-based digital cameras and video systems are also used to perform “plate rounds” in remote hospitals and clinics within a multihospital system. Newer Internet technologies involving robotic microscopes and high-resolution video systems now allow microbiologists to change the focus and change the slide positioning of a microscope located anywhere in the world and to view the resulting images on a monitor in their office. The availability of digital photomicroscopy has significantly enhanced the microbial identification process, and it has helped to standardize microbe identification. A wide variety of microscopes can be purchased with integrated camera systems and sophisticated light metering and exposure controls. Accessory cameras are also available from a large number of aftermarket manufacturers. However, an expensive camera system does not automatically confer the ability to produce high-quality images. Publicationquality photomicrographs require proper specimen illumination and optical train alignment to achieve the microscope’s ultimate potential (5). Color photography can be especially demanding because specimens may appear yellow or blue under tungsten halogen (3,200 K) light, depending upon whether the lamp voltage is above or below the recommended 9-volt setting. Newer camera systems have sophisticated exposure, lighting, and white balance controls that make image capture easier, but they cannot correct for poor technique. Not all microbiologists can afford a microscope with an integrated camera system. Simple eyepiece cameras can be used to capture bright-field images for Internet consults, training manuals, and plate rounds. The simplest configuration for eyepiece photography involves the use of a pointand-shoot digital camera. A number of adapters that allow

12 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

coupling a fixed-lens camera to a microscope eyepiece tube are now available. Photographs taken in this manner are often acceptable, but they may be dark and have some chromatic aberration (due to different lens correction factors) and vignetting (pipe view effect). Another method for photomicroscopy is to use the camera port on microscopes fitted with a trinocular head. Olympus and Nikon have introduced adapters that allow their digital cameras to attach to the camera tube of their microscopes. In addition, camera tube and eyepiece adaptors for a number of digital cameras are available from Microscope Depot (Tracy, CA). Photography under these conditions is best done using a camera with through-the-lens exposure metering. These devices work well if the exposure is not longer than several seconds or shorter than 1/3 second (5). Many of these cameras have built-in flashes that should be turned off during photomicroscopy. Consumer-grade digital cameras may have problems with fluorescence microscopy due to the extreme contrast of fluorescent images and the tendency of metering systems to average exposure values over the entire field (5).

CARE AND USE OF THE MICROSCOPE Proper care and maintenance of the microscope will prolong the usable life of the instrument and allow for more accurate interpretation of microbiological images. The microscope should be kept in a low-vibration, lowdust environment to facilitate viewing and to decrease damage to the optical systems. The optical elements should be kept completely free of dust, dirt, oil, solvents, and other contaminants (23). Ideally, the microscope should be covered and the lamp should be turned off when the microscope is not in use. Do not touch the optical surfaces with your fingers (23). Keep the lenses clean, and be sure to remove oil or mounting fluid from the objectives, condenser, and mechanical stage after each session. Immersion oils act as a slow-acting solvent that can weaken the optical mounting cement (7). Avoid dragging the high dry objective through oil or fluorescence mounting fluid. One way to avoid accidental contact with these fluids is to place the high dry objective and the oil immersion objective in the nosepiece on opposite sides of the low-power objective (5). Lenses should be dusted with a fine lens brush and a bulb syringe and cleaned with lens paper and a commercial lens cleaner that is approved by the microscope manufacturer (7). Compressed air is not recommended because it can leave a residue on glass surfaces (7). Commercial cleaners, such as Windex and Sparkle, should not be used on optical surfaces because they often contain acids or bases that can erode the antireflection coatings on the lens (7). Organic solvents, such as alcohols and acetone, should not be used on the lenses because these solvents may dissolve the optical mounting cement (7). Unused spaces in the nosepiece (Fig. 3) should be plugged, and the eyepieces should remain installed at all times to prevent introduction of dust into the body tube. The stage should be cleaned regularly, and any spilled immersion oil or mounting fluid must be removed, or slides will stick when they are moved across the stage. Oils and mounting fluids also collect dust and grit, which can damage the optical and mechanical parts. Microscopists should not attempt to remove or disassemble the objectives, as this increases the potential for damage (23). This is a job that is best left to professionals (23). Do not use lubricating oils on the gears or bearing surfaces of the microscope because this may cause

the condenser and stage to sink from their own weight (23). Annual or semiannual cleaning and adjustment by a professional microscope repair person will also help to extend the usable life of the microscope.

ERGONOMICS Peering into a microscope eyepiece for long periods is not an activity for which the body is well adapted. Microscope work requires the head and arms to be locked in a forward position and inclined toward the microscope with rounded shoulders. This unusual positioning is further exaggerated when the feet are placed on the ring-style footrests that are common on many laboratory stools. Poor posture and awkward positioning during microscopy can cause pain or injury to the neck, wrists, back, shoulders, and arms (24). In one regional survey of cytotechnologists, Kalavar and Hunting (25) found that 70.5% of respondents reported neck, shoulder, or upper back pain during microscopy and that 56% had an increased prevalence of hand/wrist symptoms. Eyestrain, leg discomfort, and foot discomfort have also been documented with long-term microscope use (26). With older microscopes, users often have their heads inclined up to 45 degrees from vertical and their upper backs may be inclined by as much as 30 degrees. Even 30-degree inclinations of the head can produce significant muscle contractions, fatigue, and pain (26). For this reason, microscopists should be taught to sit upright and hold their head in a neutral position (27). During microscopy, the laboratorian should sit erect while maintaining the natural curve of the spine (27). The lower back and shoulder blades should be supported by the chair, and a lumbar support cushion should be used if necessary. The legs and feet should rest firmly on the floor or a footrest. The chair should have a pneumatic height adjustment (23), and the seat should have a sloping front edge to prevent undue pressure on the thighs. The backrest should be adjustable for both height and angle. The chair should have a five-pointed star base with caster wheels. Knee spaces, which are often used for laboratory storage, should be free from obstructions, and there should be a minimum of 2 inches of clearance between the thigh and the bottom of the desk or counter (25). Obstructions that prevent the microscopist from holding his or her shoulders perpendicular to the ocular axis of the microscope should be removed (24). The upper arms should be perpendicular to the floor with the elbows close to the body. The forearms should be parallel with the floor, and the wrists should be straight. The head should be upright, and the neck should bend as little as possible, preferably no more than 10 to 15 degrees. The eyepieces should be just below the eyes, and the eyes should look downward at a 30- to 45-degree angle. The use of tilting microscope heads can significantly improve the comfort of the microscopist (25, 28, 29). Repetitive motions of the hands and the contact stress of arms resting on (the edge of) a hard surface can cause pain and nerve injury, leading to repetitive stress injuries and/ or carpal tunnel syndrome (24). The use of padded armrests can moderate some of these problems. In addition, microscopes should not be placed under an air vent in order to prevent stiffening of the muscles during microscopy. Most laboratory microscopes are used by multiple individuals, and it is often a challenge to find conditions or microscope configurations that satisfy everyone. Some laboratories place microscopes on books or heavy blocks of wood

2. Microscopy n

to accommodate taller microscopists (23). This configuration creates a number of problems. If the microscope is raised to a sufficient height to prevent neck flexion, users may be forced to bend their wrists into an unnatural position. If the microscope is lowered to allow the forearms to remain parallel to the floor, the neck is forced to bend. Lowering the chair to its lowest position causes leg discomfort. Shorter individuals may have to raise the chair to a level where their feet no longer touch the floor. Footrests can ameliorate this problem, but some individuals may have insufficient space under the benchtop to accommodate their legs. In practice, most laboratories will elect to use a suboptimum, but workable, microscope configuration that all users can employ. Under these conditions, microscopists can reduce stress and fatigue by taking 1-min “microbreaks” every 10 to 15 min during which they can stand, stretch, and allow the eyes to focus at a distance. Eye fatigue can be a major problem for microscope users, especially if they have poor vision. The diopter adjustment provided on most microscope eyepieces can be adjusted to compensate for minor near- and far-sightedness and allow the user to remove his/her glasses during microscope use. The diopter adjustments do not adjust for astigmatism, and users with moderate to severe astigmatism should wear glasses when using the microscope. Most microscope manufacturers now produce high-eyepoint eyepieces that move the visual observation point further from the eyepiece, thereby facilitating the use of glasses during microscopy. Ensuring that the microscope images are as bright, sharp, and crisp as possible will also help to reduce eye fatigue and associated headaches. The importance of proper alignment of the microscope and optical components cannot be overstressed. Proper optical alignment and the use of newer objectives with higher NA values will produce brighter images and better resolution, which eases the strain of searching for tiny specimen details. The use of a neutral blue (daylight) filter during bright-field microscopy can also help to lessen eye strain when examining microbiological specimens. In the future, many new microscopes will display the specimen image on a computer monitor. This innovation may alleviate many of the eyestrain problems that develop during extended microscope use (29). Microscopes are as different as the people who use them, and the previous comments should not be construed as a prescription for alleviating strain or repetitive-motion injuries in every situation. When purchasing a microscope, every effort should be made to allow microscopists to evaluate the new microscope under their normal working conditions. Some microscopes will be comfortable for some users and uncomfortable for others. In the long run, the fit and feel of the microscope is just as important as the optical characteristics.

CONCLUSIONS Advances in the design, resolution, and ergonomics of modern microscopes have greatly enhanced our ability to study and identify microorganisms. Microscopy still has a central role in the detection of infectious agents despite highly publicized advances in DNA and RNA detection systems. Microscopic examination of clinical specimens provides a rapid and inexpensive “first pass” in the detection and identification of infectious agents. Thus, clinical microscopy will continue to be a core competency in clinical microbiology laboratories for the foreseeable future.

13

REFERENCES 1. Douglas SD. 1985. Microscopy, p. 8–13. In Lennette EH, Balows A, Hausler WJ Jr, Shadomy HJ (ed), Manual of Clinical Microbiology, 4th ed. American Society for Microbiology, Washington, DC. 2. Abramowitz M. 2003. Microscope Basics and Beyond, vol 1. Olympus America, Inc, Melville, NY. 3. Abramowitz M. 1988. Contrast Methods in Microscopy: Transmitted Light, vol 2. Olympus America, Inc, Melville, NY. 4. Abramowitz M. 1994. Optics, a Primer. Olympus America, Inc, Melville, NY. 5. Abramowitz M. 1998. Photomicrography, a Practical Guide, vol 5. Olympus America, Inc, Melville, NY. 6. Delost MD. 1997. Introduction to Diagnostic Microbiology. Text and Workbook, p 37–41. Mosby-Year Book, Inc, St Louis, MO. 7. Murphy DB, Davidson MW. 2013. Fundamentals of Light Microscopy and Electronic Imaging. Wiley-Blackwell, Hoboken, NJ. 8. Abramowitz M. 1993. Fluorescence Microscopy: The Essentials, vol 4. Olympus America, Inc, Melville, NY. 9. Coons AH, Creech HJ, Jones RN. 1941. Immunological properties of an antibody containing a fluorescent group. Exp Biol Med 47:200–202. 10. Coons AH, Creech HJ, Jones RN, Berliner E. 1942. The demonstration of a pneumococcal antigen in tissues by use of fluorescent antibody. J Immunol 45:159–170. 11. Coons AH, Kaplan MM. 1950. Localization of antigen in tissue cells, II. Improvements in a method for the detection of antigen by means of fluorescent antibody. J Exp Med 91:1–13. 12. Gardner PS, McQuillin J. 1974. Rapid Virus Diagnosis: Application of Immunofluorescence, 2nd ed. Butterworth, London, United Kingdom. 13. Goldman M. 1968. Fluorescent Antibody Methods. Academic Press, New York, NY. 14. Johnson GD, Nogueira Araujo GM. 1981. A simple method of reducing the fading of immunofluorescence during microscopy. J Immunol Methods 43:349–350. 15. Nairn RC. 1976. Fluorescent Protein Tracing, 4th ed. Livingstone, London, United Kingdom. 16. Riggs JL, Seiwald RJ, Burckhalter J, Downs CM, Metcalf TG. 1958. Isothiocyanate compounds as fluorescent labeling agents for immune serum. Am J Pathol 34:1081–1524. 17. Xing Y, Chaudry Q, Shen C, Kong KY, Zhau HE, Chung LW, Petros JA, O’Regan RM, Yezhelyev MV, Simons JW, Wong MD, Nie S. 2007. Bioconjugated quantum dots for multiplexed and quantitative immunohistochemistry. Nat Protoc 2:1152–1165. 18. Kairdolf BA, Smith AM, Stokes TH, Wang MD, Young AN, Nie S. 2013. Semiconductor quantum dots for bioimaging and biodiagnostic applications. Ann Rev Chem 6:143–162. 19. Reid T, Baldini A, Rand TC, Ward DC. 1992. Simultaneous visualization of seven different DNA probes by in situ hybridization using combinatorial fluorescence and digital imaging microscopy. Proc Natl Acad Sci U S A 89:1388– 1392. 20. Tkachuk DC, Pinkel D, Kuo WL, Weier HU, Gray JW. 1991. Clinical applications of fluorescence in situ hybridization. Genet Anal Biomol Eng 8:67–74. 21. Johnson GD, Davidson RS, McNamee KC, Russell G, Goodwin D, Holborow EJ. 1982. Fading of immunofluorescence during microscopy: a study of the phenomenon and its remedy. J Immunol Methods 55:213–242. 22. Giloh H, Sedat JW. 1982. Fluorescence microscopy: reduced photobleaching of rhodamine and fluorescein protein conjugates by n-propylgallate. Science 217:1252–1255.

14

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

23. Murray RGE. 1999. Introduction to morphology, p 5–20. In Gerhardt P, Murray RGE, Wood WA, Kreig NR (ed), Methods for General and Molecular Bacteriology. ASM Press, Washington, DC. 24. Kofler M, Kreczy A, Gschwendtner A. 2002. “Occupational backache”—surface electromyography demonstrates the advantage of an ergonomic versus a standard microscope workstation. Eur J Appl Physiol 86:492–497. 25. Kalavar SS, Hunting KL. 1996. Musculoskeletal symptoms among cytotechnologists. Lab Med 27:765–769.

26. Chaffin D, Andersson G. 1991. Occupational Biomechanics. John Wiley & Sons, Inc, New York, NY. 27. Thompson SK, Mason E, Dukes S. 2003. Ergonomics and cytotechnologists: reported musculoskeletal discomfort. Diagn Cytopathol 29:364–367. 28. Kofler M, Kreczy A, Gschwendtner A. 1999. Underestimated health hazard: proposal for an ergonomic microscope workstation. Lancet 354:1701–1702. 29. Vratney M. 1999. Considerations in microscope design to avoid cumulative trauma disorder in clinical laboratory applications. Am Clin Lab 18:8.

Laboratory Detection of Bacteremia and Fungemia MICHAEL L. WILSON, MELVIN P. WEINSTEIN, AND L. BARTH RELLER

3 specimens may be mishandled, leading to falsely negative results; (iv) some microorganisms cannot be recovered adequately using broth-based blood cultures; (v) the number of microorganisms in a given specimen may be too low to be recovered by current methods; and (vi) current use of 4- or 5-day incubation and testing cycles on automated systems will result in a small number of isolates being undetected. It was long expected that molecular methods would provide a gold standard test for the detection of bacteremia or fungemia, but this has not been the case. As will be reviewed below, some molecular methods are not even as sensitive as cultures, some have been shown to detect nucleic acids in persons without bacteremia or fungemia, and some proteins in blood have been shown to act as inhibitors to some nucleic acid amplification assays. One approach has been to use composite standards, such as the use of combined results from different assays. From a statistical standpoint, this is a flawed analysis because any given assay cannot be compared against itself (as part of the gold standard). Moreover, the entire point of developing a standard is to be able to use the standard in many different studies; these “combined” gold standard assays preclude that for the obvious reason that it quickly becomes impractical to use multiple assays as a standard in every evaluation. Assays also change over time, or are no longer marketed at all, which would eliminate their use as part of a combined gold standard. For all these reasons, and despite their shortcomings, blood cultures remain the imperfect gold standard laboratory test for the diagnosis of bacteremia and fungemia.

The laboratory detection of bacteremia and fungemia remains one of the most important—and complex—roles of clinical microbiology laboratories. This is, in part, because the attributable mortality for bacteremia and fungemia remains as high as 12% (1) but also because rapid, accurate, and reliable identification of patients with bacteremia or fungemia is of critical importance in influencing treatment. Blood culture results guide antimicrobial therapy and also subsequent surgical procedures, removal of vascular access lines, and other clinical interventions. Moreover, diagnosis of bacteremia and fungemia requires more than just a single test. It requires more than one blood culture, drawn from different sites, identification of isolates recovered from blood specimens, antimicrobial susceptibility testing of isolates, and interpretation of results in conjunction with other tests and cultures. The goal of this chapter is to provide a summary of the clinical importance of bacteremia and fungemia, the scientific and medical principles underlying current diagnostic methods, a summary of alternative diagnostic approaches, and a brief review of those tests that are emerging as potential additions to or replacements for traditional procedures.

ASSESSMENT OF METHODS FOR DETECTING MICROORGANISMS IN BLOOD No method has been shown to be an ultimate standard for the detection of either bacteria or fungi from blood. It has been recognized for decades that even with the best blood culture systems, use of optimal methods for collecting blood specimens, and limiting collection of blood specimens from patients with a high pretest probability of bacteremia or fungemia, usually only 8 to 12% of blood cultures will yield microbial isolates, of which one-third to one-half will be contaminating skin flora. The reasons for this are not fully understood. The single biggest cause is that many blood cultures are obtained from patients who are at low or no risk for bacteremia or fungemia. A recent study, using a Bayesian prediction model of objective clinical and laboratory risk factors, showed the likelihood of true bacteremia to range from 0.4 to 18.4%, with a mean prevalence of 6.9% in a large cohort of patients (2). Other causes also play a role: (i) many patients are receiving empiric antimicrobial therapy at the time blood specimens are collected, thereby reducing the yield from blood cultures; (ii) patients may have temporarily cleared microorganisms from blood; (iii)

DIAGNOSTIC IMPORTANCE Determining which patients have bacteremia or fungemia and subsequent identification of pathogens and their antimicrobial susceptibility profiles are the most important objectives for using blood cultures as a diagnostic test. However, the identity of pathogens and the pattern of recovery from blood cultures provide important diagnostic clues as to the location and type of infection. There are strong associations between sites of infection and which pathogens are recovered from the bloodstream (3), observations that give critically important information to providers as to the nature of the infection. It should be remembered that up to 29% of blood culture isolates do not have an identifiable source of infection (1).

doi:10.1128/9781555817381.ch3

15

16

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

PROGNOSTIC IMPORTANCE The attributable mortality rate of about 12% is an overall rate for hospitalized adult patients with bacteremia or fungemia. When the site of infection, type of service, and other variables are used to further categorize patients, mortality rates have been shown to vary widely (1, 3). There is less known about the prognostic importance of blood culture isolates recovered from outpatients as a group, primarily due to a lack of controlled studies but also because most patients with bacteremia or fungemia are sick enough to be admitted to the hospital and therefore become inpatients. Moreover, the frequency of community-acquired occult bacteremia in children has been greatly reduced or eliminated where conjugate pneumococcal and Haemophilus influenzae type b vaccines are used. What data have been published indicate that, for bacteremic patients with uncomplicated pyelonephritis or other uncomplicated infections that do not require subsequent hospitalization, blood cultures generally are not needed because the results have minimal prognostic (and little or no diagnostic) value (4–7).

and to limit the overall number of blood draws to those that are essential for patient care. For older children, the volume of blood that should be drawn for culture differs little or not at all from that specified for adults. There are no controlled clinical trials that provide guidance, but broadly speaking, children age 12 and older (except for very small children) should have blood cultures drawn according to the criteria used for adult patients. For younger children, the total volume of blood that should be drawn for culture is lower than that for adult patients. For neonates, infants, and very young children, the volume of blood drawn for culture needs to be reduced by two mechanisms: the volume drawn per culture as well as the number of cultures. The volume of blood drawn should not exceed 1% of the patient’s blood volume, but this may need to be modified if the patient is anemic at the time of the blood draw (8). As discussed in the following section, the number of blood cultures drawn at one time is critical for proper interpretation of blood culture results. Therefore, no fewer than two blood cultures should be drawn from pediatric patients.

Number of Blood Cultures

CRITICAL FACTORS During the past 40 years, a number of studies established and clarified which factors were most important in the recovery of pathogens from patients with bacteremia or fungemia. Many of the findings are pertinent today, while others now largely have historic interest only. The factors that remain the most important are described in the following paragraphs, but it should be emphasized that optimal recovery of pathogens from blood requires that all of these factors be addressed together, not in isolation.

Volume of Blood Cultured For adult patients and older children, the volume of blood cultured is the most important factor in recovery of pathogenic microorganisms from blood. It is still important in younger children, but there are practical limits to the volumes of blood that can be withdrawn for laboratory tests such as blood cultures (8). This importance stems from the observation that there is a direct relationship between the total volume of blood cultured and the likelihood of recovery of pathogens (8–12). The research that established this often was done in conjunction with controlled clinical evaluations of different blood culture bottles and systems, which guided development of many of the commercial products that are available today. The outcome of many studies was that, for adult patients, 8 to 10 ml of blood should be inoculated into each of two bottles per blood culture, for a total of 16 to 20 ml of blood. This observation, combined with the finding that 2 to 4 blood cultures are needed to detect bacteremia or fungemia during a septic episode, means that up to 80 ml (four blood cultures of 20 ml each) should be drawn to optimize microbial recovery. Although this is a large amount of blood, and the full 80 ml is not necessary for all patients, it may be necessary to establish whether or not a patient is bacteremic or fungemic depending on the clinical situation and causative organism. This volume of blood may be reduced by half to 40 ml with use of blood culture bottles containing certain additives. For patients with anemia or other reasons for concern about the overall volume of blood drawn for laboratory testing, it is of no benefit to decrease the volume of blood drawn per culture; to do so will only result in decreased recovery of pathogens. A better diagnostic alternative is to modify the number of blood cultures drawn (as discussed below)

Collecting the correct number of blood cultures has two benefits. First, it helps ensure that an adequate volume of blood is drawn for culture. Second, it allows providers to correctly interpret results of blood cultures. Single blood cultures yield information that is difficult to interpret, unless the isolate is one that rarely, if ever, is recovered as a contaminant (e.g., pathogenic fungi such as Histoplasma capsulatum). Aside from examples such as that, however, most isolates recovered from blood cultures require the isolation from more than one blood culture to be considered a cause of sepsis. Contaminants generally occur in only one blood culture of a series, whereas pathogens typically occur in more than one of the blood cultures in a series. For patients with intravascular foci of infection, such as infective endocarditis, all blood cultures in a series should yield the pathogen. A common question concerns the diagnostic sensitivity and specificity of a blood culture. Because the clinical presentation of patients with bacteremia or fungemia is so varied, clinical signs and symptoms cannot be used as a gold standard against which blood cultures can be prepared. There also is no other laboratory test (or combination of tests) that can serve as a surrogate gold standard. In the absence of another gold standard test, then, only estimates can be made regarding diagnostic sensitivity and specificity. Four studies have addressed this question directly (9,13– 15). Because these studies were conducted during the period from 1975 to 2007, only rough comparisons should be made of the results. This is because (i) the studies used different blood culture systems; (ii) different volumes of blood were cultured; (iii) the relative distribution of pathogenic species changed during this time; (iv) markedly different classes of antimicrobial agents were in use; (v) this time period was the one in which a large number of intravenous devices, implants, and prostheses were introduced and later used widely; (vi) new pathogens were discovered; and (vii) the study designs all were different. A summary of the results from these four studies is shown in Table 1. The question, then, is whether the traditional recommendation of drawing two to three blood cultures for the detection of common pathogenic bacteria and yeasts remains valid. Clearly, a single blood culture is insufficient. Because two blood cultures will only recover 80 to 90% of pathogens, at least three blood cultures are necessary, as

3. Laboratory Detection of Bacteremia and Fungemia n TABLE 1 Cumulative percentage of recovery of pathogenic microorganisms by number of blood cultures No. of cultures

% Recovery of microorganisms in study by: Washingtona

Weinstein et al.b

Cockerill et al.c

Lee et al.d

80 88 99

91.5 >99

67.4 81.8 95.7 100

73.1 85.7 98.2 99.8

17

relative effects these four properties have on recovery of pathogenic microorganisms from blood cultures is unknown; however, controlled clinical trials have shown improved recovery of Gram-negative rods and streptococci with adequate concentrations of SPS.

Agitation 1 2 3 4

a Data from reference 13. Of the four groups in this table, this is the only set that excluded patients with infective endocarditis. b Data from reference 3. c Data from reference 9. d Data from reference 15.

that will recover 96 to 98% of pathogens. Although four blood cultures will recover close to 100% of pathogens, routinely drawing this number of blood cultures has several drawbacks: (i) it is not necessary for all patients, (ii) it adds substantially to costs if done on all patients, and (iii) it isn’t practicable for some patients. One recommended approach is to draw two blood cultures in the first 24 h, followed by an additional two blood cultures during the subsequent 24 h (9). The main criticism of this approach is that it can delay detection of bacteremia for an additional 24 h. Because this approach also suffers from the aforementioned drawbacks, the available evidence supports drawing three blood cultures in the first 24 h. Although adult patients vary substantially in both height and weight, in general the number of blood cultures drawn does not need to be adjusted by either parameter for previously healthy adult patients. For patients with anemia, for whom withdrawing blood for laboratory testing may worsen their anemia, the most important decision is whether blood cultures (or other laboratory tests) are necessary at all. If blood cultures are clinically indicated, it makes neither clinical nor laboratory sense to decrease the number of blood cultures or volume drawn; to do either is not in the patient’s best interest.

Dilution of Blood Dilution of blood was more important with older blood culture media, where the antimicrobial effects of blood per se, and antimicrobial agents in the blood, needed to be diluted by higher volumes of broth medium. Over time it became clear that, with modern blood culture systems using standardized blood culture media, anticoagulants, and other factors, a blood-to-broth ratio of between 1:5 and 1:10 was sufficient. With the addition of additives to the broth medium that bind or sequester antimicrobial agents, a bloodto-broth ratio of as little as 1:4 may be sufficient.

Anticoagulants In the past, a number of different anticoagulants have been considered for blood culture bottles, since clotted blood diminishes yield (16). Almost all have now been abandoned based on bacterial inhibition studies with seeded blood cultures. Most commercial blood culture bottles today contain sodium polyanethol sulfonate (SPS) in a narrow range of concentrations; some also have sodium citrate alone or in combination with SPS. SPS, in addition to its anticoagulant property, inhibits complement activity, inactivates clinically achievable concentrations of some aminoglycoside antibiotics, inactivates lysozyme, and blocks phagocytosis. The

Agitation of aerobic blood culture bottles, by any mechanism, has been shown to increase recovery of pathogens, and all automated systems include agitation of aerobic bottles and most anaerobic bottles as well.

Medium and Additives Although at one time there were many types of broth media used for blood cultures, today most commercial blood culture bottles contain soybean casein digest broth, also known as Trypticase soy broth (BD). Other types of media have shown comparatively little advantage over soybean casein digest broth, even those that were designed to recover specific groups of pathogens. In the past, a wide variety of additives were added to broth media, many of which are no longer available. Current additives, which were designed to improve recovery of pathogens primarily from patients receiving antimicrobial therapy at the time of culture, clearly are beneficial in terms of recovery of pathogens and have the added advantage of allowing for use of smaller volumes of blood without a concomitant decrease in microbial recovery. Use of some additives may also result in increased recovery of bacterial contaminants.

SPECIMEN COLLECTION Skin Disinfection Trials of different skin disinfectants have been published for half a century, with development of reasonable conclusions as to which disinfectants perform better. It cannot be overemphasized, however, that the margin of difference between disinfectants is small and that what is by far more important is the technique used to disinfect skin (17). As a general guide, the best skin disinfectant is chlorhexidine, followed by tincture of iodine, povidone-iodine, and then various alcohols. Specific procedures for disinfection of skin are provided with each product, and so will not be presented here, but should be followed closely by users. For infants less than 2 months of age, chlorhexidine should not be used but rather alcohol swabs should be used. Perhaps the single most important point to be made is that skin disinfection takes time: time to perform the procedure and adequate time for the disinfecting solution to work. As noted below, adequacy of disinfection can easily be audited by monitoring contamination rates. For most adults and older children, blood should be drawn from veins in the antecubital fossae. Not only are these veins readily accessible by venipuncture, but also they are sufficiently large to allow for drawing blood by a needle and syringe (i.e., a “butterfly” apparatus is not needed) and the veins are less likely to collapse. Although the practice is common, as a best practice, blood cultures should not be drawn through indwelling vascular access lines, owing to higher contamination rates. For patients with suspected line-related infection, blood should be drawn through the line and from venipuncture sites and the results compared (8, 18), as described below.

18

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

Number and Timing of Cultures As stated previously, three to four blood cultures should be drawn simultaneously to detect bacteremia or fungemia. A single blood culture lacks sufficient sensitivity to detect bacteremia, and for many isolates, the result cannot be interpreted (e.g., coagulase-negative staphylococci). There is no benefit to waiting an arbitrary amount of time between drawing blood cultures (19), even though that practice was widely used for decades. For patients with suspected infective endocarditis, or other intravascular foci of infection, three microbiological factors are of importance: documenting bacteremia, determining the identity of the infecting pathogen, and characterizing the antimicrobial susceptibility profile. Documenting continuous bacteremia, while necessary in some cases, is probably of less relative importance for most patients. In cases where it is necessary because of a confusing clinical or microbiological presentation, drawing three to four blood cultures spaced 30 to 60 min apart is a reasonable approach. For patients with prosthetic valves or an implanted device, for whom infections are likely to be caused by coagulase-negative staphylococci, skin disinfection is of critical importance to minimize the risk of contamination. It is also important to not draw blood cultures through indwelling vascular catheters in these patients. Most cases of infective endocarditis are caused by common bacterial pathogens, and therefore, blood cultures do not require use of special media or prolonged incubation and monitoring. For patients who are more likely to have infection caused by Abiotrophia spp. or Granulicatella adiacens, although special media are not necessary, it is important to notify the laboratory so that the pattern of growth on agar plates can be monitored (e.g., lack of growth on sheep blood agar plates) (8). The duration of incubation does not need to be more than 5 days; two studies have documented that all isolates from patients with infective endocarditis are recovered within that time (9, 20).

INTERPRETATION OF BLOOD CULTURE RESULTS Unlike straightforward laboratory tests that yield a binary test result of positive or negative, blood cultures yield a complex set of data that can only be interpreted in the context of the clinical scenario that prompted their collection. The information generated by blood cultures includes the identity of any bacteria or fungi isolated from the specimen (or combination of microorganisms), the number of positive and negative cultures, and the time for detection of microbial growth, all of which must be interpreted in light of the pretest probability for bacteremia or fungemia; the results of other clinical, laboratory, or radiographic findings; type(s) of isolates recovered from other body sites; response (or lack of response) to therapy; clinical signs and symptoms; and the clinical judgment of the ordering physician. In routine clinical practice, interpretation should not rely solely on the identity of the microorganism(s) isolated. While some isolates almost never are contaminants (e.g., Brucella spp., Francisella tularensis, and Histoplasma capsulatum) and are easy to interpret, isolation of most bacteria and fungi requires further interpretation. Isolates such as Staphylococcus aureus, Streptococcus pneumoniae, Escherichia coli, other members of the family Enterobacteriaceae, Pseudomonas aeruginosa, and Candida albicans are usually

pathogens but, in a minority of cases, can represent contaminants (3). In contrast, isolation of many other species (e.g., Propionibacterium spp., Bacillus spp. other than Bacillus anthracis, and most Corynebacterium spp.) almost always represents contaminating flora. Determining the clinical importance (if any) is often difficult for the coagulasenegative staphylococci, as the identity of isolates per se is of little or no help. This is particularly true for patients with indwelling catheters, implants, or prosthetic devices, for whom the coagulase-negative staphylococci are one of the most common causes of infection. Another good example is isolation of Staphylococcus lugdunensis, which can cause infections of foreign devices or of host tissue in a manner similar to that of S. aureus, but it also can be a contaminant (21). Isolation of this bacterium from blood cultures, as with other coagulase-negative staphylococci, requires additional information for proper clinical interpretation. Isolation from even a single culture can be clinically important. Interpretation of blood culture results collected by venipuncture is more straightforward than that for blood cultures collected from indwelling venous lines. For blood cultures collected via lines, interpretation involves analysis of the additional factors of the type of line, where it is inserted in the body, how long it has been in place, and whether isolates are recovered from the line only, from an accompanying blood culture collected via venipuncture, or from both. A number of interpretive criteria have been proposed, but the multiplicity of studies with various methodologies makes interpretation and comparison difficult. Although complex by necessity, the current CLSI criteria (8) remain useful and are shown in Table 2. Other criteria have been suggested for making the interpretation of blood culture results more accurate and reliable. The time required for an automated blood culture system to flag a bottle as positive (i.e., to detect a growth signal), or so-called “time to positivity,” has been studied in a number of settings. Although there is a correlation between the earlier detection of pathogens compared with that of contaminants, as documented many years ago, there is so much overlap between the time needed to detect growth of both groups that the information is of minimal clinical use. Moreover, time to positivity obviously is not useful for slowly growing microorganisms and probably is not useful in patients who are receiving antimicrobial therapy, which can delay detection of microbial growth. One exception to this principle is that most isolates recovered after more than 72 h of incubation represent contaminants, with the obvious exceptions of some slowly growing pathogenic bacteria and yeasts. Another commonly used criterion is that of considering the number of positive bottles, rather than the number of positive cultures. Although there are conflicting data regarding this issue, the most thorough study, by Mirrett and colleagues (22), showed that, for coagulase-negative staphylococci, there is no correlation between the number of bottles positive in blood culture sets and the clinical importance of the isolates. This observation held true for sets that consisted of two, three, or four blood culture bottles (22). Fewer data are available regarding other types of blood culture isolates, but because microorganisms in blood cultures show a Poisson distribution, there is no reason that the principle for coagulase-negative staphylococci should not hold true for other types of isolates. Last, the issue of quantitative blood cultures still arises on occasion. Although methods for quantitation (or semiquantitation) have been devised and there are some data to

3. Laboratory Detection of Bacteremia and Fungemia n TABLE 2

19

Interpretive criteria for CRBSIa, b

Short-term peripheralc catheters Method Obtain two sets of peripheral blood cultures via venipuncture Remove catheter aseptically and culture using semiquantitative method of Maki Interpretation

If one or more peripheral blood culture sets are positive AND the catheter segment culture is positive (≥15 colonies) for the same microorganism: suggestive of CRBSI If one or more blood peripheral culture sets are positive AND the catheter segment culture is negative: inconclusive (except for Staphylococcus aureus or Candida spp.) If both peripheral blood culture sets are negative AND the catheter segment culture is positive (any colony count): suggestive of catheter colonization If both peripheral blood culture sets are negative AND the catheter-segment is negative: CRBSI is unlikely Nontunneled and tunneled central venous catheters and VAPd Method Obtain two sets of peripheral blood cultures with at least one set drawn via venipuncture; the other set should be drawn aseptically from the catheter hub or through the VAP septum Interpretation If both sets are positive for the same microorganism: suggestive of CRBSI If both sets are positive for the same microorganism AND the set drawn through the catheter becomes positive ≥120 min earlier: suggestive of CRBSI If both sets are positive for the same microorganism AND the set drawn through the catheter becomes positive ≤120 min earlier: CRBSI is still possible if both sets yield the same microorganism with an identical antimicrobial susceptibility profile If both sets are positive AND the set drawn through the catheter has at least a 5-fold-greater colony count than the peripheral culture (for laboratories using a manual quantitative method such as lysis-centrifugation): suggestive of CRSBI If only the blood culture set drawn from the catheter becomes positive: inconclusive for CRBSI and suggests either colonization of the catheter or contamination during specimen collection If only the blood culture set drawn peripherally becomes positive: inconclusive for CRBSI except if S. aureus or Candida spp. is isolated; in those cases, documentation of CRBSI requires isolation of the same microorganism by culture of the catheter tip segment or additional positive catheter or peripheral blood cultures Alternative method Obtain two sets of peripheral blood cultures via venipuncture Remove catheter aseptically and culture by semiquantitative method of Maki Interpretation If one or both of the peripheral blood culture sets AND the catheter segment culture are positive with the same microorganism: CRBSI is likely If one or both of the peripheral blood culture sets are positive AND the catheter-segment culture is negative: this may represent CRBSI if positive for S. aureus or Candida spp.; in those cases, documentation of CRBSI requires further isolation of the same microorganism by additional peripheral blood cultures If the peripheral blood culture sets are negative AND the catheter segment culture is positive: suggests colonization If the peripheral blood culture sets and the catheter segment culture are negative: CRBSI is unlikely a

For all categories, interpretations are in the absence of any other identifiable source of infection. CRBSI, catheter-related bloodstream infections. Text modified from reference 8. The term “peripheral” in this table refers to blood specimens obtained via venipuncture of a peripheral vein. d VAP, venous access ports. b c

support this approach, in reality, the approach is not practicable for routine use. Moreover, as newer technologies are developed with increasing analytical sensitivity, it is possible that methods will be devised to quantitate the number of pathogens in blood via surrogate markers.

LABORATORY DETECTION— CULTURE-BASED METHODS Conventional detection of bacteremia or fungemia has been either direct, via recovery of pathogenic microorganisms by culture of blood specimens, or indirect, by identification of surrogate markers. The latter method was investigated widely during the 1970s and 1980s by attempts to identify methods such as detection of limulus endotoxin assays, or detection of bacterial antigens in blood, but these efforts are largely only of historic interest. As a result, culture

remains the only widely used laboratory method for detection of bacteremia or fungemia. Blood cultures were first described more than a century ago and, by the 1930s, had prompted a number of investigations that defined some of the methods still used today. By the 1960s, blood cultures were widely used, but methods varied considerably, a multiplicity of methods were in use (with varying degrees of success), and it became increasingly clear through time that the overall approach left much to be desired. During the 1970s, John Washington and other investigators began a series of experiments and controlled clinical trials to define the critical factors in the detection of bacteremia and fungemia, identify best practices for blood cultures, and establish scientific evidence upon which future blood culture systems could be developed and used on a widespread basis. Out of these studies emerged a growing body of knowledge regarding the critical factors for recovering pathogenic microorganisms from blood, knowledge that

20

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

has resulted in more refined clinical approaches to blood cultures and to development of commercially available blood cultures systems that incorporate these critical factors into their design and use.

Manual Blood Culture Systems Relatively few nonautomated blood culture systems are still manufactured and sold commercially in the United States. Those that are available commercially are well characterized as to their diagnostic strengths and limitations. These systems are adequate for the detection of common bacterial and fungal pathogens, are easy to use, require only a nonCO2 incubator, and are inexpensive (12). The main drawback to manual systems is that they are labor-intensive, which precludes their use in laboratories processing even moderate numbers of blood cultures. Moreover, with the availability of smaller instrumented blood culture systems, even laboratories processing low numbers of blood cultures can opt for using an automated system. The first manual blood culture systems consisted of little more than blood culture bottles containing various medium formulations, an anticoagulant, and occasionally additives or supplements. Detection of microbial growth was via two methods: subcultures after 24 or 48 h and at the end of the incubation period (so-called terminal subcultures) and visual observation of bottles for hemolysis (color change), gas production, or turbidity. This approach was not practicable for laboratories performing more than a small number of blood cultures. Because early instrumented/automated blood culture systems had their own issues with practicability, a number of alternative manual blood culture systems were developed and marketed. Of these, only two remain available in most markets: Septi-Chek (BD Biosciences, Sparks, MD) and Oxoid (Remel, Lenexa, KS). Septi-Chek is a variation of the biphasic system originally developed by Castaneda to recover Brucella spp. from blood. These systems are called biphasic because there is the traditional liquid broth medium phase combined with a solid phase consisting of various agar media. In biphasic systems, blood is inoculated into the culture bottle and mixed with the broth medium. By one of several devices, the bloodbroth mixture is then flooded over the solid media that acts as a direct subculture. The Septi-Chek variant of a biphasic system consists of a plastic paddle that is attached to the bottle after receipt in the laboratory. This obviously is an aerobic system, and for recovery of anaerobic bacteria, there is a separate bottle that does not have a corresponding paddle. Bottles are then incubated (with or without agitation) and inspected visually once or twice each day for evidence of microbial growth. The agar-coated paddle not only provides a second means of detecting microbial growth but also provides for early isolation of microbial colonies. Aerobic bottles are inverted following each inspection, providing further subcultures of the blood-broth mixture. Current versions of Septi-Chek bottles contain one of several broth media with 0.05% SPS as an anticoagulant. Two bottle sizes are available: bottles containing 70 ml of broth, accommodating blood specimens of up to 10 ml (to maintain an adequate blood-to-broth ratio), and bottles containing 20 ml of broth accommodating blood specimens of up to 3 ml. Bottles are available that contain soybeancasein digest broth, soybean-casein digest/Columbia broth, soybean-casein digest/thioglycolate broth, or brain heart infusion broth. Agar paddles use chocolate, MacConkey, and malt agars. In controlled clinical evaluations, Septi-Chek has been shown to perform well and to be an acceptable manual

system for routine practice (23–27). It is not practicable for use in laboratories processing large numbers of blood cultures, but cost-effectiveness studies defining the practical upper limit of Septi-Chek bottles that can be handled before labor costs become prohibitive have not be performed. In contrast to Septi-Chek, the manual blood culture system Oxoid Signal is a one-bottle system without an anaerobic version. Oxoid is another variant of the Castaneda bottle. With Oxoid Signal, bottles are inoculated with blood through a rubber septum in a lid. The Signal device consists of an empty plastic cylinder attached to a needle, which when inserted into the bottle extends to below the surface of the blood-broth mixture. During microbial growth, release of gases into the blood-broth mixture increases the concentrations of those gases, which reach equilibrium with those in the bottle headspace. Through time, this increases the atmospheric pressure within the bottle, eventually forcing some of the blood-broth mixture upward through the needle into the attached Signal device. Although this provides a second mechanism for detecting microbial growth, unlike Septi-Chek it does not provide for earlier isolation of microbial colonies. In controlled clinical evaluations, the Oxoid Signal system has not performed as well as other blood culture systems (28–31).

Lysis-Centrifugation Only one commercial lysis-centrifugation blood culture system has been marketed, the Isolator blood culture system (Wampole Laboratories, Cranbury, NJ). With lysis-centrifugation, the blood specimen is inoculated into tubes containing a mixture of the lysing agent saponin, an anticoagulant, and a fluorocarbon-cushioning agent. The blood is then lysed by the saponin, and the mixture is centrifuged to separate the components. The supernatant is then removed and discarded, and the suspended pellet is used to inoculate whatever culture media are deemed necessary to recover specific pathogens. The system has been used for detection of routine bacterial pathogens, but because delayed processing has been reported to reduce recovery of anaerobic bacteria, some Haemophilus species, and Streptococcus pneumoniae, other commercial broth-based systems are likely better for routine use (32–36). Although at one time Isolator was a good method for recovery of pathogenic yeasts, dimorphic fungi, mycobacteria, and Bartonella spp., other systems (see below) are at least equal for recovering these pathogens. Detection of Bartonella spp. is best achieved by use of nucleic acid amplification; if not available, serologic testing can be used. These issues, combined with the manual nature of lysis-centrifugation, have made it less practicable for routine use.

Automated Blood Culture Systems As the number of blood cultures performed began to increase during the 1960s and 1970s, with the resulting need for methods that were less labor-intensive, there developed a growing interest in development of automated blood culture systems. A number of these were developed and tested, but only a few have been commercially successful. The first of these was the Bactec 460 radiometric system (Becton Dickinson, Sparks, MD), a derivative of the original Bactec 220 system that was developed in the late 1960s and first marketed in the early 1970s. During the next two decades, the Bactec 660, 730, and 860 systems in turn succeeded the 460 system. All of these systems had in common the detection of microbial growth by monitoring CO2 production by growing microorganisms. The 220 and 460 systems used radiometric detection of 14C-labeled CO2,

3. Laboratory Detection of Bacteremia and Fungemia n 21

whereas the 660, 730, and 860 systems detected CO2 production by infrared spectrophotometry. Because both radiometric and nonradiometric detection methods required removal of an aliquot of the atmosphere in the bottle headspace, monitoring more than once or twice per day was not possible. This is because an equal volume of sterile gas had to be added to bottles after sampling to maintain the appropriate pressure in each bottle. This, in turn, reduced slightly the CO2 present in the atmosphere, thereby limiting the frequency of sampling. Because once or twice daily sampling did not allow for detection of microbial growth at the earliest possible time, a newer generation of blood culture technology was needed. The next generation, conceived in the late 1980s and introduced in the 1990s, was that of the continuous-monitoring blood culture systems (CMBCS). These systems take their name from the fact that, unlike previous automated blood culture systems, which monitored bottles for CO2 production only a few times each day, the newer systems monitor CO2 production much more frequently, typically once every 10 to 15 min (12). Unlike older automated systems, which flagged potential microbial growth when the CO2 level in a bottle exceeded an arbitrary threshold level, CMBCS use one of several computer algorithms to detect microbial growth. The first is use of a threshold level, second is a sustained linear increase in CO2 production, and third is an increase in the rate of CO2 production (although specific computer algorithms used by each manufacturer are proprietary). Because the latter two criteria depend upon actively growing microorganisms, delayed placement of bottles in instruments, which allows microorganisms to grow and thereby produce CO2 prior to testing, can result in delayed (or lack of) detection of microorganisms in bottles. Because of the large amount of data to be analyzed for each bottle, and for the many bottles in each incubator, the success of CMBCS is as much due to the development of more powerful computer processors and memory as it is to anything else. For CMBCS to monitor CO2 levels on a frequent basis, two other barriers to testing had to be removed. The first was to eliminate the need for sampling and replenishing the atmosphere in the bottle headspace. This was achieved by the development of sensors that could be read through the wall of the bottles without need for invasive testing. The second was to eliminate the need for repeated manual loading and loading of bottles onto instruments for testing. Although this was achieved to some degree by the Bactec 860 system, each bottle still had to be moved to the testing apparatus. In contrast, CMBCS have a mechanism for monitoring the sensor of each bottle individually, thereby eliminating the need to ever move bottles during incubation and testing. Overall, the change to more frequent testing with CMBCS reduced the time to detect microbial growth by 1 to 1.5 days compared with Bactec radiometric and nonradiometric systems (12).

BacT/Alert The first of the CMBCS was BacT/Alert, with prototypes under development in 1988 and the first limited clinical trials conducted by 1990 (37). BacT/Alert is a colorimetric system, detecting changes in CO2 concentration in bottles via changes in color of a pH-sensitive device in the base of bottles. Since it was introduced, the system has undergone several changes in style and configuration, with the BacT/ Alert 3D being the most current version of the instrument. Sequential medium formulations include standard aerobic and anaerobic media; aerobic, anaerobic, and pediatric (aer-

obic only) fastidious antimicrobial neutralization (FAN) media (which include activated charcoal and Fuller’s earth [Ecosorb] designed to inactivate or bind antimicrobial agents); FAN aerobic (FA), FAN anaerobic (FN), and pediatric FAN (PF) bottles; FA Plus, FN Plus, and PF Plus media (which contain proprietary adsorbent polymeric beads); and the Mycobacteria Process (MP) bottle designed to recover mycobacteria. Since it was introduced more than 20 years ago, a number of clinical evaluations of BacT/Alert have been published that have established the performance characteristics of the system and different bottle types that have been produced. These findings are summarized in Table 3. Broadly speaking, these characteristics hold true for all of the bottles available with CMBCS: bottles with additives outperform bottles without additives, only a 4- to 5-day incubation and testing cycle is necessary, blind and/or terminal subcultures are not necessary, and bottles containing additives increase recovery of contaminating microbial flora.

Bactec 9000 Series In 1992, Becton Dickinson introduced the second of the CMBCS, the Bactec 9000 series. Initially there were two versions, the Bactec 9120 (holding 120 bottles per cabinet) and the Bactec 9240 (holding 240 bottles per cabinet). Several years later, the Bactec 9050 (holding 50 bottles) was introduced for laboratories needing an automated system with a smaller capacity (38). The Bactec technology is similar to that of BacT/Alert, the main difference being that Bactec CMBCS use a fluorescence-sensing mechanism to detect CO2 production. Bactec systems make use of several medium formulations, including standard aerobic/F and anaerobic/F bottles, Plus aerobic/F and Plus anaerobic/F bottles, which contain antibiotic-binding resins attached to tiny glass beads, lytic/10 anaerobic/F medium bottles, Peds Plus/F bottles, and Myco/ F lytic bottles that are designed to increase recovery of fungi and mycobacteria (39). Each of these bottle types accepts up to 10 ml of blood, except for the Peds Plus/F bottles, which accept 1 to 3 ml of blood. A large number of clinical evaluations of Bactec Standard and Plus/F bottles used on the 9050, 9120, and 9240 systems have been published. The performance characteristics of these systems are summarized in Table 4. In contrast to Standard/F and Plus/F bottles, Bactec Myco/F lytic bottles were developed to allow for recovery of mycobacteria and fungi as well as common bacterial pathogens. Myco/F lytic bottles differ from all other Bactec 9000 bottles in that the fluorescent sensor in the bottle detects decreasing oxygen concentration (i.e., detects oxygen consumption) as opposed to detecting increased concentrations of carbon dioxide. In a number of clinical trials, Myco/F lytic bottles have been shown to compare favorably with other systems for detection of mycobacteria and fungi. In an early study, the Myco/F lytic bottle detected fewer H. capsulatum isolates but more Cryptococcus neoformans isolates than the Isolator system (40). Myco/F bottles have been shown to be equivalent to ESP II bottles for recovery of mycobacteria overall, with significantly better recovery of Mycobacterium avium complex in Myco/F lytic bottles (40). More recent studies have shown Myco/F lytic bottles and BacT/Alert MB bottles to be equivalent to but appreciably faster than the Isolator 10 system for recovery of Mycobacterium tuberculosis and Mycobacterium avium complex clinical isolates (41, 42).

VersaTREK Blood Culture System The third CMBCS introduced during the 1990s was the ESP system (Difco Laboratories), now marketed by Thermo

22

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

TABLE 3

Summary of performance characteristics of BacT/Alert blood culture system

BacT/Alert standard bottles are equivalent to other commercial bottles without additives for recovery of common pathogenic bacteria and yeasts (38, 66, 67) BacT/Alert aerobic FAN bottles are superior to BacT/Alert aerobic standard bottles for recovery of common pathogenic bacteria and yeasts (68) BacT/Alert anaerobic FAN bottles are superior to BacT/Alert anaerobic standard bottles for recovery of common pathogenic bacteria and yeasts with the exception of strict aerobes such as Gram-negative nonfermenters and yeasts (69) BacT/Alert FA Plus and FN Plus bottles are superior to FA and FN bottles for recovery of common pathogenic bacteria and yeasts with earlier time to detection (70) BacT/Alert PF Plus bottles are superior to PF bottles for recovery of common pathogenic microorganisms with earlier time to detection (71) BacT/Alert FA Plus and FN Plus bottles are superior to BacT/Alert standard bottles for recovery of common pathogenic bacteria and yeasts with earlier time to detection for FA Plus bottles (70, 72) BacT/Alert FAN bottles are equivalent to other commercial bottles with additives for recovery of common pathogenic bacteria and yeasts (73, 74) BacT/Alert FAN bottles are equivalent to other commercial bottles for improving recovery of pathogens from patients who are receiving antimicrobial therapy when cultures are drawn BacT/Alert FAN bottles are superior to BacT/Alert standard bottles for detection of episodes of bacteremia and fungemia (68, 69, 75) BacT/Alert FAN bottles recover more contaminating bacteria than do BacT/Alert standard bottles BacT/Alert bottles can be processed using a 5-day rather than a 7-day incubation period without a significant decrease in recovery of pathogenic bacteria and fungi (76, 77) Terminal subcultures of bottles shown to be negative by the instrument do not increase recovery of pathogenic bacteria or yeasts BacT/Alert aerobic standard bottle inoculated with 10 ml of blood detects more microorganisms than the same bottle inoculated with 5 ml of blood (78)

Scientific (Cleveland, OH) as VersaTREK. VersaTREK differs from the BacT/Alert and Bactec systems in several fundamental ways. First, VersaTREK uses a different system for detecting microbial growth. Once received in the laboratory, bottles are fitted with an adapter mechanism before being loaded into the instrument. The detector mechanism allows for the system to monitor pressure changes within the headspace of each bottle as oxygen, hydrogen, nitrogen, and carbon dioxide are consumed or produced by growing microorganisms. Second, the blood broth mixture within aerobic bottles is agitated by use of a stir bar contained within each bottle. Anaerobic bottles are not agitated. Third, the broth medium in VersaTREK Redox 1 aerobic bottles is soy casein peptone broth, whereas the broth medium in Redox 2 anaerobic bottles is proteose-peptone

broth. Fourth, bottles are monitored for growth less frequently: at 12 min for aerobic bottles and 24 min for anaerobic bottles. Last, unlike BacT/Alert FAN and Bactec Plus bottles, both of which contain additives to minimize or negate antimicrobial activity in blood specimens, VersaTREK Redox 1 and Redox 2 bottles contain 80 ml of broth medium, the larger volumes providing greater dilution of blood and any antimicrobial agents contained therein. The smaller Redox 1 EZ Draw and Redox 2 EZ Draw bottles, which are marketed to allow for direct collection of blood specimens into the bottles, still contain 40 ml of broth medium for greater dilution of blood specimens compared with BacT/Alert and Bactec bottles. A number of clinical evaluations of VersaTREK have been published. Based on the results of these evaluations, the performance characteristics of this system are summarized in Table 5.

TABLE 4 Summary of performance characteristics of Bactec blood culture systems

Pediatric Blood Culture Bottles

Bactec standard bottles are equivalent to other commercial bottles without additives for recovery of common pathogenic bacteria and yeasts (79) Bactec Plus/F bottles are superior to BacT/Alert standard bottles for recovery of common pathogenic bacteria and yeasts (80) Bactec anaerobic Plus/F bottles are superior to Bactec standard anaerobic bottles for recovery of common pathogenic bacteria and yeasts (80) Bactec Plus/F bottles are equivalent to other commercial bottles with additives for recovery of common pathogenic bacteria and yeasts (74, 81) Bactec Plus/F bottles are equivalent to other commercial bottles with additives for improving recovery of pathogens from patients receiving antimicrobial therapy when cultures are drawn Bactec Plus/F bottles are superior to bottles without additives for detection of episodes of bacteremia and fungemia (80) Bactec Plus/F bottles recover more contaminating bacteria than do Bactec standard bottles

A number of blood culture bottles intended for use with pediatric patients have been developed and marketed. Because of the practical difficulties in conducting clinical trials involving large numbers of hospitalized children, these products have not been evaluated to the same extent as those for adult patients. While there is no drawback to using these bottles (other than cost differences), there appears to be little advantage to using them in regions where immunization with pneumococcal conjugate and Haemophilus influenzae type b vaccines are given widely.

LABORATORY DETECTION— NON-CULTURE-BASED METHODS Surrogate Markers for Sepsis A number of nonmicrobiological tests have been evaluated for their ability to detect bacteremia or fungemia. Although most of these evaluations state that assays were evaluated for the ability to detect sepsis, it should be remembered that sepsis is foremost a clinical diagnosis. What most of these

3. Laboratory Detection of Bacteremia and Fungemia n TABLE 5 Summary of performance characteristics of VersaTREK blood culture system VersaTREK bottles are superior to Bactec standard bottles for recovery of common pathogenic bacteria and yeasts A number of studies have shown VersaTREK bottles to be inferior for recovery of Staphylococcus aureus compared with bottles from other systems containing additives (33, 73, 82, 83), whereas one study found the opposite (84) Several studies have reported decreased recovery of contaminants from VersaTREK bottles (11); another study found the opposite (85) Several studies have reported increased recovery of streptococci and enterococci from VersaTREK bottles (82, 83, 86); another study found the opposite (84) In some studies, VersaTREK bottles have been reported to detect significantly more pathogens in blood taken from patients receiving antimicrobial therapy at the time of blood culture (82, 83, 86) VersaTREK bottles can be processed using a 4- or 5-day rather than a 7-day incubation period without a significant decrease in recovery of pathogenic bacteria and fungi (87, 88)

studies were evaluating, in fact, was the ability to detect patients with bacteremia or fungemia who subsequently developed the clinical signs and symptoms of sepsis. Early assays, such as the limulus amoebocyte assay, showed promise but were never fully developed or marketed. Perhaps the most widely studied assays have been those that measure procalcitonin and serum lactate levels. For procalcitonin, the available evidence shows that the assay is not useful for the detection of bacteremia or fungemia, due to a lack of both sufficient sensitivity and specificity. The assay has been shown to be useful, however, when used to monitor patients with a diagnosis of bacteremia or fungemia as a guide to stopping antimicrobial therapy. The findings with serum lactate levels are similar. The assay lacks both sensitivity and specificity for the diagnosis of bacteremia or fungemia, but serial measurements may be of use both for prognosis and for modifying antimicrobial therapy.

Nucleic Acid Amplification Methods A number of laboratory-developed assays designed for the direct detection of bacteremia or fungemia from blood specimens have been evaluated and reported (43). As presented elsewhere in this chapter, although laboratory-developed assays may serve a role as a proof of concept, they are of little use to the community at large because the assays cannot be marketed or distributed to other laboratories. Moreover, development and use of such assays is likely prohibitively expensive and beyond the expertise available in most clinical laboratories. SeptiFast (Rotkreuz, Switzerland) is currently available in parts of the European Union but not in the United States. The system is a real-time PCR assay designed to detect, directly from blood, 10 species of Gram-negative bacilli, 6 species of Gram-positive cocci, 5 species of yeasts, and 1 filamentous fungus. The method uses the LightCycler system; although this technology is available for use in the United States, SeptiFast reagents are not. Overall, the system has shown mixed results in clinical evaluations (44). Taken together, the data from these evaluations indicate that SeptiFast shows comparatively low sensitivity for detecting pathogens in blood. The number of studies, conflicting study designs and definition, and other

23

factors make interpretation of the data difficult, but overall, the assay lacks sufficient sensitivity to be used as a standalone test. SeptiFast has been evaluated as an adjunct to blood cultures, in which a positive result from SeptiFast between days 3 and 7 following a positive blood culture result predicted an increased risk of patients developing complicated bloodstream infections (45). Whether the same risk stratification could be achieved using other laboratory tests has not been evaluated. Another molecular assay, Xpert MTB/RIF (Cepheid, Sunnyvale, CA), has been evaluated for detection of Mycobacterium tuberculosis directly from blood specimens. In one small clinical evaluation, the calculated sensitivity of Xpert was only 21% (a figure notably similar to some of the data for SeptiFast, as described above), but in this group of patients, a positive Xpert test result was useful for stratifying patients into those who were or were not at increased risk for early death (46). At this time, therefore, the data are clear that no molecular assay is available that can act as a substitute for blood cultures, but these tests may be useful as adjuncts to blood cultures.

RAPID IDENTIFICATION OF MICROBIAL ISOLATES A number of rapid identification methods have been introduced over the years, with increasingly effective results. Early attempts were largely modifications of biochemical tests or early molecular methods (e.g., DNA probes) that did not have the desired effect. More-recent methods have yielded substantially better results for rapid identification of select groups of microbial pathogens. Moreover, there is evidence that earlier detection and identification of bloodstream pathogens, or antimicrobial resistance, improves outcomes (47) and facilitates antimicrobial stewardship (48, 49).

Peptide Nucleic Acid-Fluorescent In Situ Hybridization The peptide nucleic acid-fluorescent in situ hybridization (PNA-FISH) method (AdvanDx, Woburn, MA), although limited to a few select pathogens, has been shown to reduce the time needed to identify microbial pathogens present in blood cultures. The principle of the method is simple, being based on the widely used principle of in situ hybridization. Unlike DNA and RNA probes, which have negatively charged sugar-phosphate backbone structures, the PNA backbone structure is composed of an electrically neutral polyamide (peptide) structure. The neutral electrical charge allows for more-rapid, tighter, and more-specific hybridization with nucleic acid targets. The probes are labeled with fluorescent dyes, which can be observed by using a fluorescent microscope. Once a blood culture bottle is flagged as positive, a smear is made of the blood-broth mixture and a Gram stain is performed. Based on those results, the appropriate PNA-FISH probe(s) can be selected for testing. The method is easy to use, only modestly expensive, and requires little in the way of infrastructure beyond that already present in most clinical microbiology laboratories (other than a fluorescent microscope) (50). However, to take full advantage of the method, it is necessary to perform the test on an ad hoc basis for individual isolates, which is not practicable in many microbiology laboratories. The method does not provide any data regarding antimicrobial susceptibility or resistance.

24

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

Nucleic Acid Amplification Tests Although nucleic acid amplification test technology would appear to lend itself to this application, surprisingly few commercial assays have been developed. In contrast, a large number of laboratory-developed assays have been reported, but their usefulness is limited by that of all user-developed assays: assays may not be adaptable for use in other laboratories, development and validation costs may be prohibitively expensive, and regulatory issues limit use of these assays in some countries (43). Their usefulness at this time is also limited by the relatively small number of pathogens that can be detected, preliminary evidence indicating that the assays are not as useful when more than one type of microorganism is in the blood-broth mixture (polymicrobial isolates), and the ability to test for only a small number of antimicrobial resistance determinants. Thus, the number of commercial assays available for this purpose is limited, as are clinical evaluations as to their effectiveness.

Multiplex Assays Two commercial systems have been developed, the Verigene assay (Nanosphere, Northbrook, IL) and the Film Array (Biofire, Salt Lake City, UT). The Food and Drug Administration has cleared both for marketing in the United States. For Verigene, separate assays for detection of either Grampositive or Gram-negative bacteria directly in blood cultures are FDA cleared; an assay for detection of common pathogenic yeasts is under development. The FilmArray product, called the blood culture identification panel, detects a combination of 24 Gram-positive bacteria, Gram-negative bacteria, and yeasts and three antimicrobial resistance genes in a single combined assay. Verigene is a multiplex assay based on PCR amplification of nucleic acid targets with subsequent detection by hybridization with oligonucleotides bound to nanosphere particles, followed by signal amplification using what the manufacturer calls a silver staining process. FilmArray also is a multiplex assay, with an initial nucleic acid extraction and purification step followed by PCR-based amplification in two stages. The first is a multiplexed reaction; this is followed by individual PCR amplification reactions designed to detect specific products from the first-stage amplification step. Final detection is based on use of endpoint melting curves. The Verigene Gram-positive blood culture assay, which is cleared for detection of 12 genera or species and three antimicrobial resistance genes, has been evaluated in several clinical trials. In the first, blood cultures processed on the VersaTREK blood culture system were used for the analysis. The evaluation of 203 total positive blood cultures showed 94% concordance for microbial identification and 97% concordance for detection of drug resistance for 178 monomicrobial isolates, with 92% and 96% concordance for 25 polymicrobial isolates (51). Results were available 24 to 48 h sooner than with conventional identification and susceptibility test methods. In the second study, blood cultures were processed using the BacT/Alert system (52). Results from this study of 186 blood cultures were similar to those of the first, in that for 176 monomicrobial cultures the concordance for identification was 96% for microbial identification and 99% for detecting drug resistance. Results were available 31 to 50 h sooner than with conventional methods. A third study showed overall concordance of 95% with conventional identification methods: 99% for monomicrobial isolates but only 33% for a small number of polymicrobial isolates (53). A fourth study was more specific and limited in scope, evaluating use of the assay as part of an antimicrobial

stewardship program targeting patients with enterococcal bacteremia (49). In this study of 74 patients, compared with the preintervention use of conventional identification and antimicrobial susceptibility testing, the postintervention phase when Verigene was used showed shortening of the time required for appropriate antimicrobial therapy to be given, shortened length of hospitalization, and lower hospital costs. Finally, in an evaluation of the assay in a pediatric hospital, the findings reported were similar to those of the first three studies: the assay showed 95.8% concordance with conventional methods for identification of Gram-positive bacteria (54). The assay also showed 100% correlation for detection of methicillin-resistant S. aureus isolates, 100% correlation for detection of vancomycin-resistant E. faecium isolates, and 98% detection of methicillin-resistant Staphylococcus epidermidis isolates (54). As in earlier studies, time to detection was substantially shorter with the Gram-positive blood culture assay than with conventional methods (54). The Verigene Gram-negative blood culture test, which is cleared to detect eight genera or species and six antimicrobial resistance genes, has been evaluated in only one published clinical trial (55). In this evaluation of 102 isolates, the Gram-negative blood culture test showed 97.9% concordance with conventional identification methods. The reported performance characteristics for detecting antimicrobial resistance or susceptibility were a positive predictive value of 95.8% and a negative predictive value of 100% (55). Only one clinical evaluation of FilmArray blood culture identification has been published (56). In this study, FilmArray showed 91.6% concordance with conventional identification methods for samples with monomicrobial growth and 71% concordance for polymicrobial samples. The assay did detect additional microorganisms in 3.6% of samples for which isolates were not recovered by conventional culture methods.

Other Nucleic Acid Amplification Tests The Gene Xpert (Cepheid, Sunnyvale, CA) has several assays that can be used in conjunction with blood cultures for identification of blood culture isolates or for detecting antimicrobial resistance. The two assays that would fit into this category are an assay for detecting the mecA gene in methicillin-resistant strains of Staphylococcus epidermidis and Staphylococcus aureus and one for detecting the vanA gene in vancomycin-resistant strains of enterococci. As with other methods for detecting pathogens or antimicrobial drug resistance in blood culture isolates, these assays do not replace blood cultures but are only an adjunct.

Antigen Detection The BinaxNOW Staphylococcus aureus test (Alere Scarborough, Inc., Scarborough, ME) is an immunochromatographic assay that uses polyclonal antibodies to detect an S. aureus-specific protein, thereby allowing for differentiation between S. aureus and other Gram-positive cocci in blood culture bottles. Current FDA clearance is for use with BacT/ Alert blood culture bottles. In one off-label study using VersaTREK blood culture bottles, the test showed 95.8% sensitivity and 99.6% specificity compared with culture and a direct tube coagulation test, both performed on aliquots of the blood-broth mixture (57). In a second off-label evaluation, the assay was compared with conventional methods using Bactec blood culture bottles (58). In this study, the BinaxNOW assay showed 97.6% sensitivity and 100% specificity.

3. Laboratory Detection of Bacteremia and Fungemia n

MALDI-TOF (MS) Matrix-assisted laser desorption ionization–time of flight (mass spectrometry) (MALDI-TOF [MS]) has been the most widely evaluated approach to rapid microbial identification of blood culture isolates. Currently two manufacturers of MALDI-TOF (MS) systems have instruments cleared by the FDA for marketing in the United States to identify microbial isolates from solid media (i.e., not directly from blood culture bottles): the Microflex LT Biotyper (Bruker Daltonics, Bremen, Germany) and the Vitek MS IVD (bioMérieux, Marcy l’Etoile, France). In an early study, 90% of bacterial isolates were identified directly from positive blood culture bottles (59). Other early studies of this technology also showed good results for identifying bacteria and fungi directly from blood culture broth specimens. One evaluation of the Biotyper evaluated the ability of MALDI-TOF (MS) to identify bacterial pathogens in 212 blood cultures (60). Of these, 42 (19.8%) showed insufficient numbers of bacteria in the blood-broth mixtures for MALDI-TOF (MS) to identify the bacteria (61). Of the other 170 blood cultures, MALDITOF (MS) showed 95.3% concordance in correctly identifying the bacteria compared with conventional identification methods (61). In another evaluation using the Biotyper (62), 330 positive blood culture bottles were analyzed, of which 318 showed growth on subcultures and 12 were considered to be false-positive signals by the blood culture instrument. Of the latter group, the MALDI-TOF (MS) results were fully concordant with culture results (62). For the 318 blood cultures that yielded growth on cultures, all were monomicrobial. When compared with results of conventional identification methods, MALDI-TOF (MS) results were concordant to the species level for 83.3% and to the genus level for 96.6% of blood cultures (62). In the most comprehensive comparison of the two MALDI-TOF (MS) systems, a total of 202 positive blood culture bottles processed on the Bactec system were tested with both versions of MALDI-TOF (MS) (62). In this evaluation, there were 181 monomicrobial and 21 polymicrobial isolates. Biotyper correctly identified 177 of 181 (97.8%) monomicrobial isolates compared to 167 of 181 (92.3%) identified by the Vitek system (62). Neither system performed well for identification of polymicrobial isolates (62). Although time to identification was not evaluated in this study, two previous studies showed a reduction in the time to identification of between 26.5 and 34.3 h compared to conventional methods (63, 64). Despite this, the inability to perform antimicrobial susceptibility testing limits the usefulness of this method.

DIRECT RAPID ANTIMICROBIAL SUSCEPTIBILITY TESTING FROM BLOOD CULTURE BOTTLES A large number of studies have attempted to answer whether direct testing of the blood-broth mixture (without the intervening step of subcultures) using conventional (not molecular) methods can be used to decrease the time needed for susceptibility test results. Theoretical obstacles to this approach are straightforward. The main obstacle is the inability to standardize the number of microorganisms in the blood-broth mixture to be used for testing. None of the CMBCS flag bottles as positive based on the number of microorganisms present. Depending on which system is used, the initial number of microorganisms present in the blood specimen, the metabolic characteristics of the microorganism (e.g., growth rate and gas production), and the number of microorganisms per milliliter when bottles are

25

flagged as positive may vary. Second, testing aliquots of the blood mixture, with or without additives, introduces a complex liquid matrix (i.e., blood, broth, anticoagulant, and any additives) that was not intended for use in commercial antimicrobial susceptibility assays. Even with centrifugation, or other procedures, this “matrix effect” cannot be eliminated or mitigated fully. A third issue, although perhaps not as important, concerns the presence of antimicrobial agents present in blood. Because these agents are given in dosages to achieve blood concentrations at or above levels designed to inhibit bacterial growth, even with dilution by the broth medium, there may still be residual antimicrobial activity in the blood-broth mixture (65). Fourth, this is a moving target. Interpretive criteria for breakpoints may change over time, thereby requiring repeated validation of the method, which would not be practicable in most settings. A fifth obstacle is regulatory: most commercial antimicrobial susceptibility assays do not include direct testing of a blood-broth mixture in the package insert, making such use off-label and, in some cases, not reimbursable. Last, the published evidence on these approaches is not persuasive: some published studies have shown these approaches to work, but others have arrived at the opposite conclusion. In an era that emphasizes evidence-based laboratory medicine, objective analysis of the literature yields the conclusion that current evidence does not support this practice.

QUALITY AUDITS AND BENCHMARKS The most commonly studied, documented, and reported quality metric regarding blood cultures is the contamination rate. There has been a long-standing recommendation that blood culture contamination rates be kept at or below 3% for hospitalized patients. This figure is not derived from anything more than the belief that it generally is not possible to maintain rates below 1% and that rates above 5% result in a confounding of the clinician’s ability to distinguish between contaminants and pathogens. Because blood culture contaminants result in increased health care costs, contamination rates above 5% also are associated with increased costs. Whether the 3% figure is realistic for outpatient settings, particularly in emergency departments, is another unanswered question. For patient safety, quality, and costs, it makes sense to target the lowest possible contamination rates, but targeting specific rates should be done with the understanding that different contamination rates occur in different settings. Another common assessment of quality is the number of blood cultures drawn per septic episode. As noted previously, interpretation of blood culture results depends heavily upon drawing both an adequate volume of blood and more than one culture. Determining the clinical importance of isolates recovered from single blood cultures can be impossible depending on the type of isolate recovered. At the same time, collecting more blood cultures than is necessary is wasteful, contributes to phlebotomy-caused anemia, and results only in recovery of more contaminants. For both reasons, laboratories should monitor the number of blood cultures collected per septic episode. A third important measurement is the adequacy of fill of blood culture bottles. Weighing filled bottles and comparing weights against those of a known standard most readily achieve this goal. Bottles filled with inadequate volumes of blood diminish yield and should be reported to the provider, with a recommendation to recollect the blood culture. Adequacy of filling should be monitored through time and by site so that any patterns of underfilling (or overfilling) can be identified and the appropriate corrective action taken.

26

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

SUMMARY Detection of bacteremia and fungemia remains one of the most important roles of clinical microbiology laboratories. Despite the development and introduction of a number of novel technologies, the blood culture using liquid-based media still remains the only practicable approach for routine patient care. Molecular detection methods show promise, but the available methods do not have the sensitivity of blood cultures (except for a select few pathogens), provide only limited information regarding antimicrobial susceptibility testing, if used alone would not allow for retention of isolates for epidemiologic investigations, at this time are not effective for detecting polymicrobial isolates, and are more expensive to use on a routine basis. Other methods such as MALDI-TOF (MS) also can achieve similar results for rapid identification of microorganisms isolated on solid media but currently are not cleared by the FDA for identification of pathogens directly from blood cultures. Experience has shown that novel technologies rarely replace older technologies but rather serve as adjunct methods to enhance older technologies. Because the isolation of pathogens from blood serves multiple clinical roles—prognosis, guiding therapy, monitoring response to therapy, and epidemiology— any approach to the laboratory detection of bacteremia and fungemia must be able to fulfill each of these roles. It is unlikely that any new technology will replace blood cultures entirely in the foreseeable future. What already is happening is that newer technologies are being integrated with blood cultures into an algorithmic approach that takes advantage of the benefits of each method.

REFERENCES 1. Pien BC, Sundaram P, Raoof N, Costa SF, Mirrett S, Woods CW, Reller LB, Weinstein MP. 2010. The clinical and prognostic importance of positive blood cultures in adults. Am J Med 123:819–828. 2. Jin SJ, Kim M, Yoon JH, Song YG. 2013. A new statistical approach to predict bacteremia using electronic medical records. Scand J Infect Dis 45:672–680. 3. Weinstein MP, Towns ML, Quartey SM, Mirrett S, Reimer LG, Parmigiani G, Reller LB. 1997. The clinical significance of positive blood cultures in the 1990s: a prospective, comprehensive evaluation of the microbiology, epidemiology, and outcome of bacteremia and fungemia in adults. Clin Infect Dis 24:584–602. 4. McMurray BR, Wrenn KD, Wright SW. 1997. Usefulness of blood cultures in pyelonephritis. Am J Emerg Med 15:137–140. 5. Mills AM, Barros S. 2005. Are blood cultures necessary in adults with pyelonephritis? Ann Emerg Med 46:285– 287. 6. Thanassi M. 1997. Utility of urine and blood cultures in pyelonephritis. Acad Emerg Med 4:797–800. 7. Velasco M, Martínez JA, Moreno-Martínez A, Horcajada JP, Ruiz J, Barranco M, Almela M, Vila J, Mensa J. 2003. Blood cultures for women with uncomplicated acute pyelonephritis: are they necessary? Clin Infect Dis 37:1127– 1130. 8. Clinical and Laboratory Standards Institute. 2007. Principles and Practice of Blood Cultures. Document M-47. Clinical and Laboratory Standards Institute, Wayne, PA. 9. Cockerill FR, Wilson JW, Vetter EA, Goodman KM, Torgerson CA, Harmsen WS, Schleck CD, Ilstrup DM, Washington JA, Wilson WR. 2004. Optimal testing parameters for blood cultures. Clin Infect Dis 38:1724–1730. 10. Mermel LA, Maki DG. 1993. Detection of bacteremia in adults: consequences of culturing an inadequate volume of blood. Ann Intern Med 119:270–272.

11. Patel R, Vetter EA, Harmsen WS, Schleck CD, Fadel HJ, Cockerill FR III. 2011. Optimized pathogen detection with 30-compared to 20-milliliter blood culture draws. J Clin Microbiol 49:4047–4051. 12. Reimer LG, Wilson ML, Weinstein MP. 1997. Update on the detection of bacteremia and fungemia. Clin Microbiol Rev 10:444–465. 13. Washington JA. 1975. Blood cultures: principles and techniques. Mayo Clin Proc 50:91–95. 14. Weinstein MP, Reller LB, Murphy JR, Lichtenstein KA. 1983. The clinical significance of positive blood cultures: a comprehensive analysis of 500 episodes of bacteremia and fungemia in adults. I. Laboratory and epidemiologic observations. Rev Infect Dis 5:35–53. 15. Lee A, Mirrett S, Reller LB, Weinstein MP. 2007. Detection of bloodstream infections in adults: how many cultures are needed? J Clin Microbiol 45:3546–3548. 16. Rosner R. 1968. Effect of various anticoagulants and no anticoagulant on ability to isolate bacteria directly from parallel clinical blood specimens. Am J Clin Pathol 68:216– 219. 17. Wilson ML, Weinstein MP, Mirrett S, Reimer LG, Fernando C, Meredith FT, Reller LB. 2000. Comparison of iodophor and alcohol pledgets with the Medi-Flex Blood Culture Prep Kit II for preventing contamination of blood cultures. J Clin Microbiol 38:4665–4667. 18. Everts RJ, Vinson EN, Adholla PO, Reller LB. 2001. Contamination of catheter-drawn blood cultures. J Clin Microbiol 39:3393–3394. 19. Li J, Plorde JJ, Carlson LG. 1994. Effects of volume and periodicity on blood cultures. J Clin Microbiol 32:2829– 2831. 20. Baron EJ, Scott JD, Tompkins LS. 2005. Prolonged incubation and extensive subculturing do not increase recovery of clinically significant microorganisms from standard automated blood cultures. Clin Infect Dis 41:1677–1680. 21. Fadel HJ, Patel R, Vetter EA, Baddour LM. 2011. Clinical significance of a single Staphylococcus lugdunensis-positive blood culture. J Clin Microbiol 49:1697–1699. 22. Mirrett S, Weinstein MP, Reimer LG, Wilson ML, Reller LB. 2001. Relevance of the number of positive bottles in determining clinical significance of coagulase-negative staphylococci in blood cultures. J Clin Microbiol 39:3279– 3281. 23. Bryan LE. 1981. Comparison of a slide blood culture system with a supplemented peptone broth culture method. J Clin Microbiol 14:389–392. 24. Pfaller MA, Sibley TK, Westfall LM, Hoppe-Bauer JE, Keating MA, Murray PR. 1982. Clinical laboratory comparison of a slide blood culture system with a conventional broth system. J Clin Microbiol 16:525–530. 25. Weinstein MP, Reller LB, Mirrett S, Wang WLL, Alcid DV. 1985. Controlled evaluation of Trypticase soy broth in agar slide and conventional blood culture systems. J Clin Microbiol 21:626–629. 26. Weinstein MP, Reller LB, Mirrett S, Wang WLL, Alcid DV. 1985. Clinical comparison of an agar slide blood culture system with Trypticase soy broth and a conventional blood culture bottle with supplemented peptone broth. J Clin Microbiol 21:815–818. 27. Weinstein MP, Reller LB, Mirrett S, Stratton CW, Reimer LG, Wang WLL. 1986. Controlled evaluation of the agar slide and radiometric blood culture systems for the detection of bacteremia and fungemia. J Clin Microbiol 23:221–225. 28. Murray PR, Niles AC, Heeren RL, Curren MM, James LE, Hoppe-Bauer JE. 1988. Comparative evaluation of the Oxoid Signal and Roche Septi-Chek blood culture systems. J Clin Microbiol 26:2526–2530. 29. Weinstein MP, Mirrett S, Reimer LG, Reller LB. 1989. Effect of agitation and terminal subcultures on yield and

3. Laboratory Detection of Bacteremia and Fungemia n

30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

speed of detection of the Oxoid Signal blood culture system versus the Bactec radiometric system. J Clin Microbiol 27:427–430. Weinstein MP, Mirrett S, Reimer LG, Reller LB. 1990. Effect of altered headspace atmosphere on yield and speed of detection of the Oxoid Signal blood culture system versus the Bactec radiometric system. J Clin Microbiol 28:795– 797. Weinstein MP, Mirrett S, Reller LB. 1988. Comparative evaluation of the Oxoid Signal and Bactec radiometric blood culture systems for the detection of bacteremia and fungemia. J Clin Microbiol 26:962–964. Cockerill FR III, Reed GS, Hughes JG, Torgenson CA, Vetter EA, Harmsen WS, Dale JC, Roberts GD, Ilstrup DM, Henry NK. 1997. Clinical comparison of Bactec 9240 Plus Aerobic/F resin bottles and the Isolator aerobic culture system for detection of bloodstream infections. J Clin Microbiol 35:1469–1472. Cockerill FR, Torgenson CA III, Reed GS, Vetter EA, Weaver AL, Dale JC, Roberts GD, Henry NK, Ilstrup DM, Rosenblatt JE. 1996. Clinical comparison of Difco ESP, Wampole Isolator, and Becton Dickinson Septi-Chek aerobic blood culture systems. J Clin Microbiol 34:20–24. Hellinger WC, Zawley JJ, Alvarez S, Hogan SF, Harensen WS, Ilstrup DM, Cockerill FR. 1995. Clinical comparison of the Isolator and BacT/Alert aerobic blood culture systems. J Clin Microbiol 33:1787–1790. Henry NK, Grewell CM, Van Grevenhof PE, Ilstrup DM, Washington JA II. 1984. Comparison of lysis-centrifugation with a biphasic blood culture medium for the recovery of aerobic and facultatively anaerobic bacteria. J Clin Microbiol 20:413–416. Henry NK, McLimans CA, Wright AJ, Thompson RL, Wilson WR, Washington JA. 1983. Microbiological and clinical evaluation of the Isolator lysis-centrifugation blood culture tube. J Clin Microbiol 17:864–869. Kiehn TE, Wong B, Edwards FF, Armstrong D. 1983. Comparative recovery of bacteria and yeasts from lysiscentrifugation and a conventional blood culture system. J Clin Microbiol 18:300–304. Thorpe TC, Wilson ML, Turner JE, DiGuiseppi JL, Willert M, Mirrett S, Reller LB. 1990. BacT/Alert: an automated colorimetric microbial detection system. J Clin Microbiol 28:1608–1612. Murray PR, Hollick GE, Jerris RC, Wilson ML. 1998. Multicenter comparison of Bactec 9050 and Bactec 9240 blood culture systems. J Clin Microbiol 36:1601–1603. Waite RT, Woods GL. 1998. Evaluation of Bactec MYCO/ F Lytic medium for recovery of mycobacteria and fungi from blood. J Clin Microbiol 36:1176–1179. Crump JA, Tanner DC, Mirrett S, McKnight CM, Reller LB. 2003. Controlled comparison of Bactec 13A, MYCO/ F LYTIC, BacT/Alert MB, and ISOLATOR 10 systems for detection of mycobacteremia. J Clin Microbiol 41:1987– 1990. Crump JA, Morrissey AB, Ramadhani HO, Njau BN, Maro VP, Reller LB. 2011. Controlled comparison of BacT/Alert MB system, manual Myco/F Lytic procedure, and Isolator 10 system for diagnosis of Mycobacterium tuberculosis bacteremia. J Clin Microbiol 49:3054–3057. Jeng K, Gaydos CA, Blyn LB, Yang S, Won H, Matthews H, Toleno D, Hsieh YH, Carroll KC, Hardick J, Masek B, Kecojevic A, Sampath R, Peterson S, Rothman RE. 2012. Comparative analysis of two broad-range PCR assays for pathogen detection in positive-blood-culture bottles: PCR–high-resolution melting analysis versus PCR-mass spectrometry. J Clin Microbiol 50:3287–3292. Westh H, Lisby G, Breysse F, Böddinghaus B, Chomarat M, Gant V, Goglio A, Raglio A, Schuster H, Stuber F, Wissing H, Hoeft A. 2009. Multiplex real-time PCR and

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

27

blood culture for identification of bloodstream pathogens in patients with suspected sepsis. Clin Microbiol Infect 15:544– 551. Fernández-Cruz A, Marín M, Kestler M, Alcalá L, Rodriguez-Créixems M, Bouza E. 2013. The value of combining blood culture and SeptiFast data for predicting complicated bloodstream infections caused by gram-positive bacteria or Candida species. J Clin Microbiol 51:1130–1136. Feasey NA, Banada PP, Howson W, Sloan DJ, Mdolo A, Boehme C, Chipungu GA, Allain TJ, Heyderman RS, Corbett EL, Alland D. 2013. Evaluation of Xpert MTB/ RIF for detection of tuberculosis from blood samples of HIV-infected adults confirms Mycobacterium tuberculosis bacteremia as an indicator of poor prognosis. J Clin Microbiol 51:2311–2316. Barenfanger J, Graham DR, Kolluri L, Sangwan G, Lawhorn J, Drake CA, Verhulst SJ, Peterson R, Moja LB, Ertmoed MM, Moja AB, Shevlin DW, Vautrain R, Dallahon CD. 2008. Decreased mortality associated with prompt Gram staining of blood cultures. Am J Clin Pathol 130:870–876. Bauer KA, West JE, Balada-Llasat JM, Pancholi P, Stevenson KB, Goff DA. 2010. An antimicrobial stewardship program’s impact with rapid polymerase chain reaction methicillin-resistant Staphylococcus aureus/S. aureus blood culture test in patients with S. aureus bacteremia. Clin Infect Dis 51:1074–1080. Sango A, McCrater YS, Johnson D, Ferreira J, Guzman N, Jankowski CA. 2013. Stewardship approach for optimizing antimicrobial therapy through use of a rapid microarray assay on blood cultures positive for Enterococcus species. J Clin Microbiol 51:4008–4011. Stone NRH, Gorton RL, Barker K, Ramnarain P, Kibbler CC. 2013. Evaluation of PNA-FISH yeast traffic light for rapid identification of yeast directly from positive blood cultures and assessment of clinical impact. J Clin Microbiol 51:1301–1302. Samuel LP, Tibbetts RJ, Agotesku A, Fey M, Hensley R, Meier FA. 2013. Evaluation of a microarray-based assay for rapid identification of gram-positive organisms and resistance markers in positive blood cultures. J Clin Microbiol 51:1188–1192. Wojewoda CM, Sercia L, Navas M, Tuohy M, Wilson D, Hall GS, Procop GW, Richter SS. 2013. Evaluation of the Verigene gram-positive blood culture nucleic acid test for rapid detection of bacteria and resistance determinants. J Clin Microbiol 51:2072–2076. Beal SG, Clurca J, Smith G, John J, Lee F, Doern CD, Gander RM. 2013. Evaluation of the nanosphere Verigene gram-positive blood culture assay with the VersaTREK blood culture system and assessment of possible impact on selected patients. J Clin Microbiol 51:3988–3992. Mestas J, Polanco CM, Felsenstein S, Bard JD. 2014. Performance of the Verigene gram-positive blood culture assay for direct detection of gram-positive organisms and resistance markers in a pediatric hospital. J Clin Microbiol 52:283–287. Mancini N, Infurnari L, Ghidoli N, Valzano G, Clementi N, Burioni R, Clementi M. 2014. Potential impact of a microarray-based nucleic acid assay for rapid detection of gram-negative bacteria and resistance markers in positive blood cultures. J Clin Microbiol 52:1242–1245. Altun O, Almuhayawl M, Ullberg M, Ozenci V. 2013. Clinical evaluation of the FilmArray blood culture identification panel in identification of bacteria and yeasts form positive blood culture bottles. J Clin Microbiol 51:4130– 4136. Dhiman N, Trienski TL, DiPersio LP, DiPersio JR. 2013. Evaluation of the BinaxNOW Staphylococcus aureus test for rapid identification of gram-positive cocci from VersaTREK blood culture bottles. J Clin Microbiol 51:2939–2942.

28

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

58. Qian Q, Eichelberger K, Kirby JE. 2014. Rapid identification of Staphylococcus aureus directly from Bactec blood culture broth by the BinaxNOW S. aureus test. J Clin Microbiol 52:319–320. 59. Moussaoui W, Jaulhac B, Hoffmann A-M, Ludes B, Kostrzewa M, Riegel P, Prevost G. 2010. Matrix-associated laser desorption ionization time-of-flight mass spectrometry identifies 90% of bacteria directly from blood culture vials. Clin Microbiol Infect Dis 16:1631–1638. 60. Stevenson LG, Drake SK, Murray PR. 2010. Rapid identification of bacteria in positive blood culture broths by matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 48:444–447. 61. Ferreira L, Sanchez-Juanes F, Porras-Guerra I, GarciaGarcia MI, Garcia-Sanchez JE, Gonzalez-Buitrago JM, Munoz-Bellido JL. 2011. Microorganisms direct identification from blood culture by matrix-assisted laser desorption/ ionization time-of-flight mass spectrometry. Clin Microbiol Infect 17:541–551. 62. Chen JHK, Ho PL, Kwan GSW, She KKK, Siu GKH, Cheng VCC, Yuen KY, Yam WC. 2013. Direct bacterial identification in positive blood cultures by use of two commercial matrix-assisted laser desorption ionization–time of flight mass spectrometry systems. J Clin Microbiol 51:1733– 1739. 63. Lagacé-Wiens PRS, Adam HJ, Karlowsky JA, Nichol KA, Pang PF, Guenther J, Webb AA, Miller C, Alfa MJ. 2012. Identification of blood culture isolates directly from positive blood cultures by use of matrix-assisted laser desorption ionization–time of flight mass spectrometry and a commercial extraction system: analysis of performance, cost, and turnaround time. J Clin Microbiol 50:3324–3328. 64. Vlek AL, Bonten JM, Boel CH. 2012. Direct matrixassisted laser desorption ionization time-of-flight mass spectrometry improves appropriateness of antibiotic treatment of bacteremia. PLoS One 7:e32589. 65. Flayhart D, Borek AP, Wakefield T, Dick J, Carroll KC. 2007. Comparison of Bactec PLUS blood culture media to BacT/Alert FA blood culture media for detection of bacterial pathogens in samples containing therapeutic levels of antibiotics. J Clin Microbiol 45:816–821. 66. Rohner P, Pepey B, Auckenthaler R. 1995. Comparison of BacT/Alert with Signal blood culture system. J Clin Microbiol 33:313–317. 67. Wilson ML, Weinstein MP, Reimer LG, Mirrett S, Reller LB. 1992. Controlled comparison of the BacT/Alert and Bactec 660/730 nonradiometric blood culture systems. J Clin Microbiol 30:323–329. 68. Weinstein MP, Mirrett S, Reimer LG, Wilson ML, Smith-Elekes S, Chirard CR, Joho KL, Reller LB. 1995. Controlled evaluation of BacT/Alert standard aerobic and FAN aerobic blood culture bottles for detection of bacteremia and fungemia. J Clin Microbiol 33:978–981. 69. Wilson ML, Weinstein MP, Mirrett S, Reimer LG, Smith-Elekes S, Chirard CR, Reller LB. 1995. Controlled evaluation of BacT/Alert standard anaerobic and FAN anaerobic blood culture bottles for the detection of bacteremia and fungemia. J Clin Microbiol 33:2265–2270. 70. Kirn TJ, Mirrett S, Reller LB, Weinstein MP. 2014. Controlled clinical comparison of BacT/Alert FA Plus and FN Plus blood culture media with BacT/Alert FA and FN blood culture media. J Clin Microbiol 52:839–843. 71. Doern CD, Mirrett S, Halstead D, Abid J, Okada P, Reller LB. 2014. Controlled clinical comparison of new pediatric medium with adsorbent polymeric beads (PF Plus) versus charcoal-containing PF medium in the BacT/ ALERT blood culture system. J Clin Microbiol 52:1898900.

72. Lee D-H, Kim SC, Bae I-G, Koh E-H, Kim S. 2013. Clinical evaluation of BacT/Alert FA Plus and FN Plus bottles compared with standard bottles. J Clin Microbiol 51:4150–4155. 73. Kirkley BA, Easley DA, Washington JA. 1994. Controlled clinical evaluation of Isolator and ESP aerobic blood culture systems for detection of bloodstream infections. J Clin Microbiol 32:1547–1549. 74. Pohlman JK, Kirkley BA, Easley KA, Basille BA, Washington JA. 1995. Controlled clinical evaluation of Bactec Plus Aerobic/F and BacT/Alert FAN bottles for detection of bloodstream infections. J Clin Microbiol 33:2856–2858. 75. McDonald LC, Fune J, Gaido LB, Weinstein MP, Reimer LG, Flynn TM, Wilson ML, Mirrett S, Reller LB. 1996. Clinical importance of increased sensitivity of BacT/Alert FAN aerobic and anaerobic blood culture bottles. J Clin Microbiol 34:2180–2184. 76. Hardy DJ, Hulbert BB, Migneault PC. 1992. Time to detection of positive BacT/Alert blood cultures and lack of need for routine subculture of 5- to 7-day negative cultures. J Clin Microbiol 30:2743–2745. 77. Wilson ML, Mirrett S, Reller LB, Weinstein MP, Reimer LG. 1993. Recovery of clinically important microorganisms from the BacT/Alert blood culture system does not require 7 day testing. Diagn Microbiol Infect Dis 16:31–34. 78. Weinstein MP, Mirrett S, Wilson ML, Reimer LG, Reller LB. 1994. Controlled evaluation of 5 versus 10 milliliters of blood cultured in aerobic BacT/Alert blood culture bottles. J Clin Microbiol 32:2103–2106. 79. Nolte FS, Williams JM, Jerris RC, Morello JA. 1993. Multicenter clinical evaluation of a continuous monitoring blood culture system using fluorescent-sensor technology (Bactec 9240). J Clin Microbiol 31:552–557. 80. Smith JA, Bryce EA, Ngui-Yen JH, Roberts FJ. 1995. Comparison of Bactec 9240 and BacT/Alert blood culture systems in an adult hospital. J Clin Microbiol 33:1905– 1908. 81. Jorgensen JH, Mirrett S, McDonald LC, Murray PR, Weinstein MP, Fune J, Trippy CW, Masterson M, Reller LB. 1997. Controlled clinical laboratory comparison of Bactec Plus Aerobic/F resin medium with BacT/Alert Aerobic FAN medium for detection of bacteremia and fungemia. J Clin Microbiol 35:53–58. 82. Doern GV, Barton A, Rao S. 1998. Controlled comparative evaluation of BacT/Alert FAN and ESP 80A aerobic media as means for detecting bacteremia and fungemia. J Clin Microbiol 36:2686–2689. 83. Welby-Sellenriek PL, Keller DS, Ferrett RJ, Storch GA. 1997. Comparison of the BacT/Alert FAN aerobic and the Difco ESP 80A aerobic bottles for pediatric blood cultures. J Clin Microbiol 35:1166–1171. 84. Zwadyk P, Pierson CL Jr, Young C. 1994. Comparison of Difco ESP and Organon Teknika BacT/Alert continuousmonitoring blood culture systems. J Clin Microbiol 32:1273–1279. 85. Welby PL, Keller DS, Storch GA. 1995. Comparison of automated Difco ESP blood culture system with biphasic BBL Septi-Chek system for detection of bloodstream infections in pediatric patients. J Clin Microbiol 33:1084–1088. 86. Mirrett S, Hanson KE, Reller LB. 2007. Controlled clinical comparison of Versa TREK and BacT/ALERT blood culture systems. J Clin Microbiol 45:299–302. 87. Doern GV, Brueggemann AB, Dunne WM, Jenkins SG, Halstead DC, McLaughlin JC. 1997. Four-day incubation period for blood culture bottles processed with the Difco ESP blood culture system. J Clin Microbiol 35:1290–1292. 88. Han XY, Truant AL. 1999. The detection of positive blood cultures by the AccuMed ESP-384 system: the clinical significance of three-day testing. Diagn Microbiol Infect Dis 33:1–6.

Systems for Identification of Bacteria and Fungi* KAREN C. CARROLL AND ROBIN PATEL

4 Traditionally, the identification of bacteria and fungi has been based on conventional tube-based biochemical reactions, with results compared to historical charts of expected biochemical reactions. Due to the need for faster, simpler methods, manual biochemical-based testing kits and instrument-based semiautomated or automated methods were introduced. Automation in microbiology first occurred in the early 1970s with the introduction of semiautomated blood culture instruments (see chapter 3), followed by instrumented systems for identification and antimicrobial susceptibility testing of bacteria. More rapid semiautomated and automated systems for antimicrobial identification and susceptibility testing followed, relying on microorganisms’ biochemical characteristics, fatty acid patterns, and/or other metabolic properties for their identification. Commercially available biochemical platforms may include decision support software integrating identification and susceptibility test results with surveillance strategies for antimicrobial resistance and guidelines for therapy. A recently introduced automated technique for microorganism identification relies on proteomic analysis of bacterial or fungal cells using matrix-assisted laser desorption ionization–time of flight mass spectrometry (MALDI-TOF MS). Nucleic acid-based approaches, typically used for bacterial and fungal identification when biochemical and/or proteomic strategies fail, include broadrange DNA target sequencing or array-based approaches and organism-specific amplification techniques. Whether a laboratory uses manual or automated, biochemical-, protein- or DNA-based methods, the scientific approach to identify microorganisms relies on the same fundamental principles. This chapter provides an overview of technologies used for the identification of bacteria and fungi recovered from clinical specimens. Discussions relevant to systems for automated blood cultures (chapter 3), antimicrobial susceptibility testing (chapter 72), immunoassays (chapter 7), molecular diagnostics (chapter 6), and detection of Mycobacterium species (chapter 30) are found elsewhere in this Manual.

ORGANISM IDENTIFICATION SYSTEMS Overview of Methods and Mechanisms of Identification Historically, microorganisms have been identified by what we now refer to as “conventional procedures,” which include reactions in tubed media and observation of physical characteristics, such as colonial morphology and odor, coupled with results of Gram staining, agglutination tests, and antimicrobial susceptibility profiles. Over the years, identification methods miniaturized commonly used biochemical reactions into a more convenient format (http://www.asmusa .org/index.php/guidelines/sentinel-guidelines ) and developed into a system-dependent approach that became the industry standard. The system-dependent method relies on a set of substrates that are carefully selected for their positive and negative reactions. These patterns create metabolic profiles that are compared with established databases. Biochemical profiles are determined by reactions of individual organisms with each of the substrates in the system. The accuracy of the reactions is dependent upon users following the directions of the manufacturer regarding inoculum preparation, inoculum density, incubation conditions, and test interpretation. Most systems rely upon one or a combination of several indicators. These include (i) pH changes resulting from utilization of a substrate, (ii) enzymatic reactions that release a chromogenic or fluorogenic compound, (iii) tetrazolium-based indicators of metabolic activity in the presence of a variety of carbon sources, (iv) detection of volatile or nonvolatile compounds, and (v) recognition of visible growth (Table 1). Additional biochemical tests for microbial identification that use other means of detecting a positive response for a given substrate may also be included. Although no formal definition of “rapid” exists for describing the time required for results to be generated, most microbiologists expect rapid systems to provide usable results within 4 h. Clearly, the generation times of microbes (typically 30 min or longer) will not allow growth-dependent methods to generate detectable biochemical responses within this time. To overcome the problem of generation times, manufacturers of rapid systems use novel substrates with which preformed enzymes, produced by the organisms to be tested, may react to elicit responses detectable within 1 to 4 h. Proteomic analysis using MALDI-TOF MS, which enables identification of bacteria and fungi in a matter of

*This chapter contains information presented by Caroline Mohr O’Hara, Melvin P. Weinstein, and J. Michael Miller in chapter 14 of the 8th edition of this Manual, by Melvin P. Weinstein and Karen C. Carroll in chapter 15 of the 9th edition of this Manual, and by Cathy A. Petti, Melvin P. Weinstein, and Karen C. Carroll in chapter 3 of the 10th edition of this Manual.

doi:10.1128/9781555817381.ch4

29

30

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

TABLE 1

Technologies for microorganism identification

System reactivity

Need for growth

Analyte

Indicator(s) of positive result

Example(s) of system

Color change due to pH indicator; carbohydrate utilization = acid pH; protein utilization or release of nitrogencontaining products = alkaline pH Color change due to chromogen or fluorogen release when colorless complex is hydrolyzed by an appropriate enzyme

API panels, Crystal panels, Vitek cards, MicroScan conventional panels, Phoenix panels, Sensititre panels MicroScan rapid panels, IDS panels, Crystal panels, Vitek cards, Phoenix panels, Sensititre panels Biolog

pH-based reactions (mostly 15–24 h)

Yes

Carbohydrate utilization

Enzyme profile (mostly 2–4 h)

No

Preformed enzymes

Carbon source utilization

Yes

Volatile or nonvolatile compound detection

Yes

Organic products Color change as a result of metabolic activity transferring electrons to colorless tetrazolium-labeled carbon sources and converting the dye to purple Cellular fatty Chromatographic tracing based on acids detection of end products, which are then compared to a library of known patterns Various Turbidity due to growth of organism in the substrates presence of a substrate Nucleic acid Electropherogram or raw sequence of nucleotide bases

Visual detection of Yes growth DNA target sequencing No

PCR/ESI MS MALDI-TOF MS

No Yes

Nucleic acid Protein

Patterns of mass signals in a spectrum Patterns of mass signals in a spectrum

minutes, is rapidly being adopted into clinical microbiology laboratories (2). Molecular methods that amplify particular gene targets novel enough to distinguish among genera and species and automated sequencing technology are used for identification of difficult-to-identify microorganisms. Proteomic and molecular methods have expanded our knowledge of pathogenesis and have, in some cases, resolved erroneous taxonomic classifications.

System Construction Microbial identification systems are either manual or automated. Manual methods use the analytical skills of the technologists for reading and interpreting the tests, whereas automated systems offer a hands-off approach, providing technologists more time for other duties and providing the laboratory with increased standardization. For all systems, the backbone of accuracy is the strength and utility of the database. Databases are constructed using known, clinically relevant strains as well as type strains of most taxa. In some cases, before an organism is added to the database, it is evaluated to confirm its relationship to other strains in the same taxon by using the likelihood fraction. This compares the characteristics of the new strain to those of a typical culture of the same species. Unusual microorganisms or common microorganisms with atypical properties often cannot be reliably identified by commercial systems unless they are well-represented in the system’s database. The number of species included in a database may vary from just a few for some manual assays to thousands for automated instruments. For most commercial systems, database maintenance is a continuous process and software upgrades incorporating major taxonomic changes are provided by the manufacturer at regular intervals. Some systems may allow users to make minor changes at the local workstation.

MIDI

API 20C AUX panels Laboratory developed; MicroSeq, GenBank, RDP, RIDOM, SmartGene IRIDICA (Abbott) Bruker, bioMérieux, Andromas

System identifications are supported by algorithm-based decision making that is generally available through a computer. Occasionally, these identifications are compiled into a preprinted index, which is used to manually convert the organism’s profile number into identification. Bayes’s theorem, or modifications of it, is often the basis of algorithm construction from data matrices. Bayes’s theorem is one of the statistical methods that manufacturers of biochemical identification systems use to arrive at an identification of a certain taxon based on the biochemical reaction profile produced by the unknown clinical isolate (3). Bayes’s theorem considers two important issues in order to arrive at an accurate conclusion: (i) P (ti/ R) is the probability that an organism exhibiting test pattern R belongs to taxon ti, and (ii) P (R/ti) is the probability that members of taxon ti will exhibit test pattern R. Before testing, we make the assumption that an unknown isolate has an equal chance of being any taxon and that each test used to identify the isolate is independent of all other tests. In this case, Bayes’s theorem can be written as P(R/ti)

P(ti /R) =

Σ P(R/ti) i

By observing reference identification charts derived by conventional biochemical tests, we know the expected pattern of the population of taxon ti (e.g., Escherichia coli is indole positive and citrate negative). R in the formula is the test pattern composed of R1, R2, . . . Rn, where R1 is the result for test 1 and R2 is the result for test 2, etc., for a given taxon. We can then incorporate the percentages (likelihoods that ti will exhibit R1, etc.) into Bayes’s theorem to arrive at an accurate taxon.

4. Systems for Identification of Bacteria and Fungi n 31

Clinical microbiologists must not, however, become dependent upon these likelihoods and percentages when interpretive judgment would suggest an alternative taxonomic conclusion. Bacteria often tend to stretch the rules of nomenclature when isolated from clinical specimens, and they may not react as expected in a commercial system, even though a legitimate result is produced (e.g., lactose-positive Salmonella species or H2S-positive Escherichia coli). The result from the most reliable systems can be misleading. In these cases, an alternative method of identification must be used. D’Amato et al. have described how the systems use the database profiles and probability matrices to arrive at an identification of an unknown taxon (4). The manufacturers of commercial identification systems rely heavily on input from their customers. Laboratories are encouraged to communicate with product manufacturers about problems, such as unusual organism identifications that develop when a method or system is being used. Manufacturers depend on customer satisfaction, and most are willing to assist in problem solving or in projects that could add strength to their systems. These companies, like their users, are clearly interested in the highest quality of costeffective patient care.

CRITERIA FOR SELECTING INSTRUMENTED SYSTEMS When selecting a method for identification and antimicrobial susceptibility testing, the laboratorian must consider several important issues. Laboratory supervisors and managers should make such major decisions carefully and with expert consultation. The process begins by answering key questions about the needs for a new system in the context of laboratory versus patient benefits. Once these questions are answered, the next step is to begin the search for the right instrument or system to meet the needs of the laboratory and the medical staff. As a general rule, it is best not to be the first to purchase a new system without having seen in the peer-reviewed literature the results of evaluations performed by reputable clinical laboratories. The manufacturer’s representative can be asked to supply peer-reviewed articles about the ability of the system to correctly identify the range of isolates usually seen in the user’s laboratory. This phase requires demonstrations and conversations regarding space requirements, technical applications, manufacturer issues such as interface capabilities and service contracts, and personnel-related concerns such as sample preparation and throughput. It is often helpful to visit other laboratories similar to one’s own that are using the system under consideration to ask if they like the system, whether they would buy it again, how much downtime they have experienced, whether the service from the manufacturer has been acceptable, and whether the system has been mechanically reliable. The laboratory should select a system that has been fully evaluated and whose accuracy exceeds 90% in its overall ability to identify common and uncommon bacteria normally seen in that particular hospital or laboratory. The system should be able to identify commonly isolated organisms with at least 95% accuracy compared with conventional methods. The accuracy of antimicrobial susceptibility testing for combination panels is as important as the accuracy of identification, perhaps more so. Chapter 72 of this Manual discusses the issues involved in instrument susceptibility test methods. Finally, with increasing use of automation in clinical microbiology laboratories, it is important to understand

how automated identification systems fit in with other automated systems in the laboratory and with the laboratory information system (see chapter 5 of this Manual).

EVALUATING AN INSTRUMENT OR SYSTEM Validating a New Instrument or Method Several references provide useful information on the approach to evaluation, verification, and validation of kits, assays, and instruments in the clinical laboratory (5–9). When an identification system is added to the laboratory, laboratories must demonstrate that the system performs as described by the manufacturer (10, 11). Published reports by other laboratories that have evaluated the system in a sound, scientific manner provide the first level of evidence of acceptable performance (9). Next, the purchasing laboratory must provide evidence of acceptable performance of the new identification instrument by in-laboratory verification. Verification involves documentation of test accuracy in the laboratory where the instrument will be used (6). The Clinical Laboratory Improvement Amendments of 1988 (12) specify the conditions for systems placed into service. Although smaller laboratories may have fewer resources than larger laboratories for verification of the accuracy of an identification system, laboratory size has no bearing on the need to ensure the accuracy of laboratory identification methods and of the work performed by a laboratory in support of patient care. The role of verification by the purchasing laboratory ensures that personnel can use the system at performance levels of accuracy already documented by the manufacturer and published in the literature. The laboratorian should expect a level of 95% agreement with the existing system or reference method and accept, in the final analysis, no less than 90% agreement. This takes into account the fact that the new system may be more accurate than the old one. As of 1998, the Food and Drug Administration (FDA) no longer performs premarket [510(K)] evaluations to “clear” automated or manual phenotypic identification systems, nor does it receive or approve quality control protocols from these devices to meet the 1988 Clinical Laboratory Improvement Amendment requirements. Laboratorians must be aware that the identification component of the new or modified system that they are using is not cleared by the FDA because this approval is no longer required. This makes it even more important for laboratorians to search the literature for valid evaluations of their chosen instrument and to conduct their own in-house validation to make sure that the instrument meets the claims of the manufacturer regarding identification. Devices and methods incorporating probes, nucleic acid amplification and other genetic methods, MALDI-TOF MS, and the antimicrobial susceptibility test component of commercial instruments will continue to be reviewed by the FDA for clearance.

LIMITATIONS OF MICROORGANISM IDENTIFICATION SYSTEMS The databases of microbial identification systems must be revised frequently to accommodate newly named species. For example, had Cronobacter sakazakii (the yellow-pigmented variant of Enterobacter cloacae) not been added to the databases of these instruments, the clinical correlation of C. sakazakii with neonatal meningitis would likely be obscured if only E. cloacae had been reported. Laboratorians must be aware that the accuracy of a system is limited to the

32 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

claims of the manufacturer for the version of the database currently in the instrument and that the database may be outdated. The laboratory procedure manual must stipulate the action to be taken when a result is questionable either because of the unusual biochemical profile of the organism or because of the appearance of an unexpected susceptibility profile. A backup method must be used to achieve an accurate identification profile. Otherwise the isolate should be sent to a reference laboratory for analysis. Closely related species may be difficult or impossible to distinguish using certain systems; however, the inability to accurately identify all species within a genus does not always have a negative effect on patient outcome. For example, correct identification of all of the newly recognized Citrobacter species may not be possible for some of the systems. In this case, the effect on patient outcome because of the inability of a system to recognize Citrobacter werkmanii may be negligible, and a simple report of “Citrobacter species” may provide adequate data for patient management. Laboratorians should recognize when they might be handling a potential agent of bioterrorism; when such pathogens are suspected, laboratories should follow Laboratory Response Network protocols and not place these organisms on automated instruments (see chapter 14, “Biothreat Agents”). Users of automated systems should be aware of the limitations of commercial products with respect to their biopreparedness plans and substitute other tests for presumptive diagnosis per recommended guidelines (http://www.asmusa.org/ index.php/guidelines/sentinel-guidelines). Microorganisms suspected of being biothreat pathogens should be referred to a public health or reference laboratory for definitive identification. Likewise, organisms that are potentially hazardous to laboratory personnel (e.g., Brucella species) should not be placed on automated systems unless this can be done in such a fashion as to eliminate any risk of disease acquisition to laboratorians. As pathogens continue to evolve and taxonomic classifications are revised, laboratorians must pay attention to the manufacturer’s communications about products, such as letters, notices, or test exclusions regarding the accuracy of their methods, as well as the published literature describing the potential problems encountered by others using these identification systems. Likewise, the user has a responsibility to report continued problems with a system or product where poor performance may lead to adverse patient outcomes.

PHENOTYPIC IDENTIFICATION SYSTEMS Automated Instruments (Vitek, Phoenix, MicroScan, TREK) The last five decades have witnessed an evolution of sophistication in automated organism identification and antimicrobial susceptibility testing systems. There are several systems available, and this section will highlight their features with respect to organism identification. Chapter 72 will discuss their antimicrobial susceptibility testing performance. The interested reader is encouraged to seek out the latest information provided by the manufacturer, as companies are constantly updating their products and, with the advent of MALDI-TOF MS, will be enhancing expert systems to link identification by proteomics with updated and expanded susceptibility testing panels. The first automated identification system to become available for clinical laboratories more than 40 years ago was the Vitek system (bioMérieux, Inc., Durham, NC).

This system was developed by NASA to test astronauts for unusual organisms acquired during the burgeoning space expeditions. The current versions, called the Vitek 2 systems, consist of a personal computer (PC), reader/incubator, and smart carrier station. The smart carrier station is a sample preparation module that standardizes the inoculum and identifies the specimen through a barcode label prior to loading the cassettes. There are several available systems depending upon the desired instrument footprint and volume of testing needed (Table 2). Depending upon the size, the instruments can accommodate 15, 30, 60, or 120 cards (Vitek 2XL). Two reader/incubator instruments can be connected to one computer. The PC has a bidirectional interface capability with the laboratory information system (www.biomerieux-usa.com). The Vitek system has cards for the identification of anaerobes and coryneforms; yeast; Neisseria species, Haemophilus species, and other fastidious organisms; Gram-positive organisms; and Gram-negative pathogens (Enterobacteriaceae, non-Enterobacteriaceae, and highly pathogenic organisms such as Brucella and Francisella species) (Table 2). The system also contains an advanced expert system that matches 2,000 organism phenotypes with 100 resistance mechanisms. Organisms for which the phenotype and MIC values match are flagged green and do not require user verification. Isolates that flag yellow (inconsistent results), red (unknown phenotype), or purple (phenotype is not in the database) require user review and problem resolution (www.biomerieux-usa.com). The Vitek 2 system, as well as the other automated systems, continues to be reviewed in the literature as new features are added and software versions are updated. New publications revisit these systems in the context of comparisons to novel molecular array platforms and MALDI-TOF MS. A few recent papers have compared the performance of several platforms; this information is useful for laboratories seeking to purchase one of these systems (13–17). Siemens Healthcare Diagnostics, Inc. (Deerfield, IL) manufactures the MicroScan WalkAway system. The available instruments include the WalkAway-40 plus and the WalkAway-96 plus, which accommodate 40 and 96 panels, respectively. The autoSCAN-4 system is a small instrument designed for low-volume laboratories; it accommodates and reads one panel at a time within several seconds. The instruments have front panel controls, and the systems include a PC keyboard, monitor, and LabPro software. The MicroScan uses system-wide bar code authentication of panels to minimize potential errors from manual labeling and/or keystroke entry when placing panels into the instrument (www.siemens.com/diagnostics). There are a variety of MicroScan panels—conventional, rapid, specialty, and Synergies plus—which are available in over 60 configurations. The conventional panels contain traditional biochemicals for identification and broth microdilution MICs and can be read visually. The rapid panels provide organism identifications as early as 2 to 2.5 h and broth microdilution susceptibility results between 4.5 and 16 to 18 h. Results are finalized within 18 h. Synergies plus panels also provide identification within 2 to 2.5 h, and broth microdilution resistance results are flagged when ready. Specialty panels are available for identification of yeast, anaerobes, fastidious Gram-negative rods, and streptococci (www.siemens.com/diagnostics). Conventional panels contain modified biochemical and chromogenic tests for the identification of a broad range of species (Table 2) (18). Rapid panels utilize fluorogenic substrates or fluorogenic indicators to detect pH changes following substrate

4. Systems for Identification of Bacteria and Fungi n 33 TABLE 2

Features of automated systems for identification of bacteria and yeastsa

b

Manufacturer

bioMérieux

Siemens

Instruments

Vitek 2 XL Vitek 2 60 Vitek 2 Compact 60 Vitek 2 Compact 30 Vitek 2 Compact 15 MicroScan WalkAway plus

Principle(s)

Software/expert systems

119 143 26 61 52

43

8

Overnight panels turbidimetric detection of carbon source utilization; enzymatic activity Rapid panels use fluorometric detection of preformed enzymes

GN Convent. GN Rapid GN Synergies GP Convent. GP Rapid GP Synergies Yeast Anaerobe NH GP GN Streptococcus Yeast ID GP GN

116 139 139 51 53 53 42 54 20 140 161 27 64 41 137

34 36

2.5 16–18

LabPro

8–16

BDXpert, BD EpiCenter

32 31

5–18

SWIN

GEN III

TREK Diagnostic Systems Biolog

Fluorometric detection

MIDI, Inc.

Incubation (h)

GP GN NH ANC Yeast

Colorimetric and fluorometric detection

OmniLog

No. of tests

Colorimetric carbon source utilization; enzymatic activity; resistance

BD Diagnostics BD Phoenix

ARIS

Panels

Organisms in database (no. of taxa)

Carbon source utilization detection by reduction of tetrazolium violet Sherlock Cell wall fatty acid microbial analysis using gas identification chromatography system

Advanced Expert System, Observa

27 36 27 25 18 48

GP GN

2,500

95

4–24c

GP GN Yeast

1,500

NA

24c

CLIN 50 database

a Derived from: http://www.bd.com/ds/productCenter/IS.asp; www.biomerieux-usa.com; www.siemens.com/diagnostics; www.biolog.com; http://www.midi-inc.com/; http://www.trekds.com/products/sensititre/c_standardid.asp. b GP, Gram positive; GN, Gram negative; NH, Neisseria, Haemophilus; ANC, anaerobe; Convent., conventional; ID, identification; NA, not applicable. c Overnight growth on particular media is required; assay takes about 2 h to perform.

utilization and the production of specific metabolic products (19). Similar to other systems, identification is based on detection of substrate utilization, pH changes, and growth in the presence of certain antimicrobial agents. Depending upon the panel used, results may be available from 2 to 42 h after incubation at 35°C (18, 19). Results can be read manually, which is useful for resolving aberrant reactions and for smaller facilities that do not have automation. Some of the panels can be stored for up to 1 year at room temperature. The company offers a device called the PROMPT system for inoculating panels in place of having to create a 0.5 McFarland standard. The user touches three well-isolated colonies that are as large as the unique wand tip. The wand is then placed into a small bottle containing 30 ml of aqueous Pluronic-D solution. After mixing well by shaking, the suspension is poured into a seed tray for inoculation into a MicroScan panel. There is also the RENOK rehydrating inoculator that can be used to simultaneously inoculate all 96 wells of any available MicroScan panel from the seed tray created by the PROMPT device (www.siemens.com/ diagnostics). The LabPro with AlertEX contains a database of predefined rules based upon Clinical and Laboratory Standards

Institute (CLSI) recommendations (20). The user can customize the rules as needed to match formularies and infection control alerts and for other uses. The software automatically notifies the user when an isolate requires attention. This may include the need for supplemental testing, a quality control issue, or another actionable item. The epidemiology management feature can provide information to infection control and pharmacy. Antibiogram tools are based on the CLSI M39 document (18) (www.siemens.com/diagnostics). The BD Phoenix Automated Microbiology System (BD Diagnostics, Sparks, MD) became available in Europe in 2001 and in the United States in 2004. Currently, the complete system consists of the Phoenix instrument, numerous test panels, the AP instrument for panel inoculation (introduced in 2008), and the BD EpiCenter system for data management (http://www.bd.com/ds/productCenter/ IS.asp). The Phoenix instrument can test up to 99 test panels and one control panel simultaneously. There are numerous panels available for Gram-positive and Gramnegative pathogen identification (ID) and/or antimicrobial susceptibility testing (AST); a yeast identification panel was introduced in 2011. The combined ID and AST panels consist of 136 wells divided into 51 wells on the ID side

34

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

and 85 wells on the AST side. Organisms are identified on the basis of 48 individual biochemical tests using a combination of fluorometric and colorimetric detection of the various substrate reactions. Identification of Gram-negative and Gram-positive organisms requires 2 to 12 h, whereas yeast identification requires 4 to 15 h. Once results are available by the Phoenix instrument, the BDXpert system software manages the interpretation of the results. Users can finalize results directly at the Phoenix instrument, using the EpiCenter, or through the laboratory’s information system. The BD EpiCenter System is a data management software system. It has a bidirectional laboratory information system interface and tools that allow the user to create specific institutional rules for AST and other types of reporting. These tools or programs also allow the user to analyze epidemiological trends and generate reports from multiple BD instruments, not just the Phoenix instrument, including the new BD Bruker MALDI Biotyper. One of the initial problems with the workflow using the Phoenix compared to other systems was the manual set up of the panels which resulted in a longer mean time of setup than for other instruments (e.g., 3 min for Phoenix compared to 1.5 min for Vitek 2) (21). In 2008, BD introduced the AP instrument to reduce the hands-on time required to set up Phoenix panels and to better standardize the inoculum preparation. The user places ID and AST broth for up to 5 isolates into a rack that gets loaded onto the AP system for automated standardization of the ID broth inoculation and subsequent inoculation of the AST broth and addition of the AST redox indicator. The user must manually transfer the rack to an inoculation station, scan the bar code labels, pour the ID and/or AST broths into the panel, seal them, and place them into the Phoenix instrument (http://www.bd.com/ds/productCenter/IS.asp). A study by Junkins et al. reports a reduction in hands-on time by 50% using the AP instrument (22). Trek Diagnostic Systems (now part of Thermo Fisher Scientific) are best known for the extensive and customizable microbroth dilution panels for susceptibility testing (discussed in chapter 72). The company manufactures identification panels (Sensititre ID plates) that contain fewer substrates than the larger systems discussed above and are designed to identify the most commonly encountered aerobic Gram-negative and Gram-positive pathogens. The separate Gram-negative and Gram-positive plates contain 32 reaction wells containing fluorogenic substrates that allow for identification in as short a time as 5 h, and which can be extended to overnight incubation if needed. Each ID microtiter plate can test three separate organisms. ID panels can be read on the same instrumentation used to read the susceptibility MIC plates. In addition, laboratories can purchase an auto inoculator called the Sensititre AIM and a nephelometer to enhance inoculum standardization. The Sensititre ARIS 2X is a fully automated benchtop instrument that incubates and reads the bar-coded plates. The system has a 64-plate capacity and accommodates all TREK plates: MIC, breakpoint, and identification. Up to 192 tests per day can be run on a single instrument. The ARIS instrument is connected to a computer that contains the SWIN software, an expert system used for interpretation of the plates. In addition, the company has an epidemiology module on SWIN that allows the user to design five customizable reports for tracking workload, MIC values, and organisms of epidemiological significance. There are no recent publications evaluating the Sensititre Gram-negative identification panels. A publication by Staneck et al. comparing the Sensititre Gram-negative identification panel to API

20E and Rapid NFT using a large number of isolates demonstrated comparable performance for the more common genera of Enterobacteriaceae and nonenterics (23). A study from Mexico evaluated the Sensititre Gram-positive ID plate to assess its utility for identifying common staphylococci compared to API STAPH v4.1 (10). Discordant isolates were resolved by PCR and sequencing of partial sequences of the 16S rRNA, sodA, and tuf genes. The Sensititre plates correctly identified only 69% of the isolates compared to 90% by API STAPH, indicating limited utility for identification of this group of Gram-positive organisms (24). The Biolog OmniLog ID System (Biolog, Hayward, CA), introduced in 1989, is a fully automated instrument based upon the ability of an organism to utilize or oxidize a panel of 95 carbon sources. The 96-well microtiter plate uses reduction of tetrazolium violet that is incorporated into each substrate as an indication of utilization of the carbon source (25). The “carbon fingerprint” is analyzed by the software of the instrument and compared to an extensive database (GEN III) of over 2,500 species of aerobic and anaerobic bacteria, yeasts, and fungi (26). Similarity indices are used to identify the test organism as follows: similarity index of 0.75, excellent identification (25). A variety of configurations, levels of automation, and identification databases are available (26). The fully automated GEN III OmniLog ID system has the capacity to incubate and monitor up to 50 Biolog MicroPlates (26). A requirement for testing is subculture of isolates to be tested to Biolog universal growth agar (25). The Biolog system has been evaluated in the literature over the last two decades (25, 27, 28). The most recent publication evaluated the accuracy of the system for identification of “atypical” clinical isolates, that is, isolates not routinely included in routine identification databases such as certain aerobic actinomycetes, Bacillus species, and fastidious Gram-negative rods (25). In this study of 159 bacterial isolates, the OmniLog system was compared to 16S rRNA gene sequencing and an extensive panel of biochemical assays; compared to conventional methods, the overall accuracy of the Biolog system was 68.3% (25). The best performance was seen with the aerobic actinomycetes (100 versus 74% accuracy with 16S rRNA gene sequencing), while the performance for fastidious Gram-negative rods was poor (20 versus 100%) (25). At least one study has also demonstrated variable performance of the Biolog system for identification of Gram-positive cocci (27). The Sherlock microbial identification system (MIDI, Inc., Newark, DE) has been available since 1991 and spun out of a partnership between Hewlett-Packard Co. (now Agilent Technologies) and the University of Delaware’s Plant Pathology Department (www.midi-inc.com). The system can identify a broad range of bacterial pathogens (approximately 1,500 species) using gas chromatographic analysis of cellular fatty acids. The traditional method of identification requires about 1.5 to 2 h perform. Organisms to be tested are subcultured to specific media and incubated for 24 h, and then a defined biomass is transferred to tubes for fatty acid extraction before gas chromatography is performed (29). The Sherlock software identifies organisms using qualitative and quantitative pattern matching of fatty acid methyl esters (29). A more recently developed rapid system for aerobes and anaerobes called Q-FAME requires less biomass and less time (approximately 24 min), but it is not FDA cleared. The Sherlock system is used as a reference method in food microbiology, by environmental laboratories, and in some clinical laboratories. Some users have

4. Systems for Identification of Bacteria and Fungi n 35

expanded the manufacturer’s library by incorporating their own strains over time, and this has been shown to enhance identification and differentiation of genera such as Corynebacterium (30). In 2005, MIDI received FDA clearance for its aerobic bioterrorism library (http://www.midi-inc.com/). The BIOMIC V3 is a digital imaging system that automates the reading and interpretation of disk diffusion tests and the results of other commercial manual identification and AST kits or tests. This system requires off-line incubation after which the user manually places the plate or test onto the instrument. The BIOMIC is programmed to read API panels, RapID tests (Remel), and Crystal panels (Becton Dickinson). It also automates the reading of Etest MICs and broth microdilution panels (www.biomic.com).

Manual Kits, Assays, and Nonautomated Platforms An array of kits, assays, and nonautomated platforms are available for identification of a variety of common and unusual pathogens. In many circumstances, these are used to supplement automated systems for identification of fastidious organisms that may fail to adequately grow in the panels of those systems or as a backup for unusual or failed results. These assays work on the principles of turbidity due to growth of organism in the presence of a substrate. Table 3 contains the list of many of the more commonly used assays stratified by the organism groups that they identify. It is recommended that the interested reader contact manufacturers for the most up to date information regarding availability and performance.

PHENOTYPE-INDEPENDENT METHODS FOR IDENTIFICATION Phenotypic microbial identification can be limiting and is often not rapid. MALDI-TOF MS is being adopted into clinical microbiology laboratories as a rapid, cost-effective method for identification of a wide range of bacteria and fungi. For challenging organisms, nucleic acid-based methods may still be required, including broad-range DNA target sequencing or arrays and organism-specific detection techniques. Amplified DNA sequence-based methods do not necessarily require optimal growth or even a viable microorganism, enable data exchange between laboratories, and may help define taxonomic relationships between microorganisms.

PROTEOMIC IDENTIFICATION SYSTEMS: MALDI-TOF MS (BASED IN PART ON REFERENCE 31) MALDI-TOF MS provides rapid, inexpensive identification of bacterial and fungal colonies (32). MALDI stands for matrix which assists in desorption and ionization of highly abundant bacterial and fungal proteins through energy from a laser (2). Although protein extraction may be performed, the most user friendly approach is to test colonies directly by moving whole cells from a bacterial or fungal colony (using a plastic or wooden stick, loop, or pipette tip) to a “spot” on a MALDI-TOF MS target plate (a disposable or reusable plate with test spots) (Fig. 1). Spots are overlaid with matrix (or first with a formic acid solution, which is allowed to dry, and then with matrix) (33–36) and dried, and the target plate is placed into a mass spectrometer (Fig. 2). The matrix (e.g., α-cyano-4-hydroxycinnamic acid dissolved in 50% acetonitrile and 2.5% trifluoroacetic acid) isolates bacterial or fungal molecules from one another, protecting them from fragmentation and enabling their de-

sorption by laser energy; the majority of the laser energy is absorbed by the matrix, converting it to an ionized state. As a result of random collision in the gas phase, charge is transferred from matrix to microbial molecules; ionized microbial molecules are then accelerated through a positively charged electrostatic field into a time of flight, or TOF, tube. Inside the tube, which is under vacuum, the ions travel toward an ion detector, with small analytes traveling the fastest, followed by increasingly larger analytes; a mass spectrum is produced, representing the number of ions of a given mass impacting the detector over time. It is highly abundant (predominantly ribosomal) proteins which generate the mass spectrum. Although they are not individually characterized, together they provide a profile unique to individual types of microorganisms, with peaks specific to genera and species. Computer software compares the generated mass spectrum to a database of reference spectra, generating a list of the most closely related organisms with numeric rankings. Depending on how high the value (percent or score) of the top match is (and considering the next best matches), the organism is identified at the family, genus, or species level. Commercial MALDI-TOF MS systems are available from bioMérieux, Inc. (Durham, NC) and Bruker Daltonics, Inc. (Billerica, MA). Another system, called Andromas (Paris, France), is primarily used in France and will not be further discussed. AnagnosTec (Zossen, Germany) marketed a microbial database called Spectral Archiving and Microbial Identification System (SARAMIS) used with Shimadzu’s AXIMA Assurance mass spectrometer (Shimadzu, Columbia, MD), which bioMérieux acquired in 2010 and changed the name to VITEK MS RUO; bioMérieux then developed a new database, software, and algorithms called VITEK MS IVD (including a prerelease version of the VITEK MS v1.1 database, the v1 system/v1.1 database, the v2.0 system/v2.0 database and algorithms, and the v3.0 system/v3.0 database and algorithms). The FDA-cleared platform is called Vitek MS. VITEK MS Plus incorporates the VITEK MS and SARAMIS v4.0 databases. The Bruker Biotyper system includes a mass spectrometer, software, and library. The FDAcleared platform is called the MALDI Biotyper CA System. At the time of this writing, there are more published studies using the Bruker than the bioMérieux system. The systems differ in databases, identification algorithms, and instrumentation. Numeric rankings, reported on different scales, are not directly comparable. Bruker’s Microflex LT mass spectrometer is a desktop instrument, whereas bioMérieux’s is a larger floor model. Rapidly implemented progressive improvements in both systems render it difficult to compare studies because of diverse specimen preparations, organism test sets (i.e., enriched for unusual organisms or not), reference identification methods, mass spectrometers, software, interpretive guidelines, and databases. In general, the technology performs at least as well as, if not better than, automated biochemical systems for identification of common bacteria and yeast (Table 4) and better for many unusual organisms (37). Usually, organisms are either correctly identified or yield a low score/percent, indicating that identification has not been achieved; the latter typically implies no “match” in the database but can occur due to technically poor preparation. Misidentifications are unusual but occur with closely related organisms; Escherichia coli and Shigella species are notably not well-differentiated by MALDI-TOF MS. Richter et al. performed a multicenter study comparing VITEK MS v2.0 to 16S rRNA gene sequencing (with supplemental phenotypic testing as needed) for identification

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

36

TABLE 3

Summary of nonautomated identification systems for bacterial identification available in 2013a

Assay

Manufacturer

Organisms targeted

Systems for anaerobe identification API 20A bioMérieux Anaerobes BBL Crystal Anaerobe BD Anaerobes RapID ANA II Thermo Scientific (Remel) Anaerobes Rapid ID 32A bioMérieux Anaerobes

Storage No. temp(°C) oftests

Incubation

2–8 2–8 2–8 2–8

21 29 18 29

24–48 h, anaerobic 4 h, aerobic 4–6 h, aerobic 4 h, aerobic

2–8

21

18–24 h, aerobic

2–8 2–8 2–8

20 20 12

4h 24–48 h, aerobic 2 h, aerobic

2–25

30

18–20 h

2–8

29

4h

2–8

15

18–24 h

2–8

12

18–48 h

2–8

12

24–48 h

2–8

24

24–48 h

2–8

17

4h

2–8 2–8

13 19

4h 4h

Staphylococci and micrococci Staphylococci Streptococci and enterococci Gram-positive organisms

2–8 2–8 2–8 2–8

20 4 20 29

18–24 h 2 h, aerobic 4–24 h, aerobic 18–24 h

Gram-positive organisms

2–8

29

4h

Staphylococcal species Streptococci and other Grampositive cocci

2–8 2–8

18 14

4h 4h

Systems for identification of Gram-positive bacilli API Coryne bioMérieux

2–8

20

24 h, aerobic

RapID CB Plus

2–8

18

4h

Systems for Enterobacteriaceae and other Gram-negative bacilli API 20E bioMérieux Enterobacteriaceae and nonfermenting Gram-negative bacteria API Rapid 20 E bioMérieux Enterobacteriaceae API 20 NE bioMérieux Non-Enterobacteriaceae API NH bioMérieux Neisseria species, Haemophilus species, Moraxella catarrhalis BBL Crystal Enteric/ BD Diagnostics Enterobacteriaceae, some GramNonfermenter ID negative nonfermenters BBL Crystal Neisseria/ BD Diagnostics Neisseria species, Haemophilus Haemophilus ID kit species, other fastidious organisms BBL Enterotube II BD Diagnostics Enterobacteriaceae, oxidase-negative Gram-negative rods Microbact 12A, 12Eb Oxoid Enterobacteriaceae, miscellaneous Gram-negative rods Microbact 12Bb Oxoid Supplements 12A for identification of other Gram-negative rods Microbact 24Eb Oxoid Enterobacteriaceae and other Gramnegative rods RapID NF Plus Thermo Scientific (Remel) Nonfermenting and selected fermenting Gram-negative rods RapID NH Plus Thermo Scientific (Remel) Neisseria, Haemophilus, Moraxella RapID ONE Thermo Scientific (Remel) Enterobacteriaceae Systems for identification of Gram-positive cocci API Staph bioMérieux RAPIDEC Staph bioMérieux API 20 Strep bioMérieux BBL Crystal GramBD Diagnostics positive ID kit BBL Crystal rapid BD Diagnostics Gram-positive ID kit RapID Staph Plus Thermo Scientific (Remel) RapID STR Thermo Scientific (Remel)

Corynebacterium and corynebacteria-like organisms Thermo Scientific (Remel) Coryneform bacilli (40 taxa)

a

Compiled from: http://vgdusa.com/bbl-crystal-identification.htm; http://www.oxoid.com; thermoscientific.com/oxoid; http://www.biomerieux-usa.com/. The 12A and 12B are in a strip format; the 12E and 24E have microplate formats.

b

of 965 Enterobacteriaceae isolates representing 17 genera and 40 species (38). MALDI-TOF MS results agreed with the reference methods for 96.7% of isolates, with 83.8% correctly identified to the species level, 12.8% limited to genuslevel identification, and 1.7% yielding no identification. Seven isolates had wrong genus identification, including three Pantoea agglomerans isolates misidentified as Enterobacter species and single isolates of Enterobacter cancerogenus, Escherichia hermannii, Hafnia alvei, and Raoultella ornithinolytica misidentified as Klebsiella oxytoca, Citrobacter koseri, Obe-

sumbacterium proteus, and Enterobacter aerogenes, respectively. Eight isolates were misidentified at the species level. In general, the Enterobacter cloacae complex is accurately identified using MALDI-TOF MS, although some species within the complex may not be discriminated from one another (39, 40). Mass spectra of Raoultella planticola, Raoultella ornithinolytica, and Klebsiella oxytoca may be similar (41). MALDI-TOF MS can identify Campylobacter species, Arcobacter butzleri, Yersinia enterocolitica (42), and Aeromonas species (43) and differentiate Capnocytophaga canimorsus

4. Systems for Identification of Bacteria and Fungi n 37

FIGURE 1 MALDI-TOF MS workflow (from reference 2). A colony from a culture plate is placed on a “spot” on a MALDI-TOF MS target plate (a reusable or disposable plate with a number of test spots). One or many isolates is tested at a time. In this example, the Bruker Biotyper system is shown. Cells are treated with formic acid on the target plate. Following drying, the matrix is added (34). After drying of the matrix, the plate is placed into the mass spectrometer for analysis (Fig. 2). A mass spectrum is generated and compared by the system’s software against a database of mass spectra, resulting in identification of the organism (Candida parapsilosis in position A4 in the example). Reproduced by permission of Mayo Foundation for Medical Education and Research. All rights reserved. doi:10.1128/9781555817381.ch4.f1

from Capnocytophaga cynodegmi (44). Salmonella enterica subsp. enterica serovar Typhi may be distinguished from other S. enterica serovars, although typing within the genus Salmonella is generally not possible (45). It may be possible to identify Burkholderia cepacia complex members, including Burkholderia cenocepacia, Burkholderia cepacia, Burkholderia stabilis, and Burkholderia vietnamiensis (46). With database enhancement, HACEK organisms (Haemophilus, Actinobacillus/Aggregatibacter, Cardiobacterium, Eikenella, Kingella; 47) and Legionella species (48) can be identified. Francisella tularensis and Brucella species may be accurately identified; however, the Bruker Biotyper library does not contain and will not identify these organisms. Use of Bruker’s “Security Relevant” database enables their identification (49). Rychert et al. reported findings from a multicenter study evaluating the Vitek MS v2.0 system for the identification of 1,146 isolates of aerobic Gram-positive bacteria (50). For 92.8%, a single accurate, species-level identification was provided. With MALDI-TOF MS, overall identification of staphylococci (51), β-hemolytic streptococci (52), aerococci (53), and enterococci is excellent. Some α-hemolytic streptococci are problematic. S. mitis and S. oralis may be poorly differentiated from S. pneumoniae, at least with the Bruker system (54, 55); the VITEK MS system may overcome this limitation (56, 57). While MALDI-TOF MS reliably identifies viridans group streptococci to the species group level, it may not be able to discriminate some closely related species (58). Other streptococci, such as S. canis, S. dysgalactiae, and S. pyogenes and S. infantarius, S. equinus,

and S. lutetiensis may not be well differentiated from one another (33). Arcanobacterium haemolyticum and Rhodococcus equi can be reliably identified, as can all but select closely related Corynebacterium species (using lower score cutoffs than recommended by the manufacturer) (59). MALDI-TOF MS can be used to identify many clinically relevant anaerobic bacteria (60, 61). Jamal et al. reported species-level identification of 89 and 100% of 274 routinely isolated anaerobic bacteria (enriched in Bacteroides fragilis) using the Biotyper DB Update–V3.3 and the VITEK MS v1 system/v1.1 database, respectively (62). Identification of more esoteric anaerobes, including Prevotella species, has been successful in 83% of cases with user supplementation of Bruker’s Reference Library 3.2.1.0 (63). Schmitt et al. evaluated a diverse collection of 253 clinical anaerobic isolates using the Bruker system and a user-supplemented database; 92 and 71% of isolates were correctly identified to the genus and species levels, respectively (64). Barreau et al. tested 1,325 anaerobic isolates using the Bruker system and correctly identified 92.5% to the species level (using lower score cutoffs than recommended by the manufacturer) (61). Garner et al. evaluated 651 anaerobic bacterial isolates using the VITEK MS v2.0 system and reported correct species-level identification in 91.2% (65). A small number of studies have reported using MALDITOF MS for mycobacteria, an endeavor which requires special processing to kill tested bacteria (for safety), disrupt clumped cells, and break down cell envelopes. Current commercial libraries inadequately address Mycobacterium

38

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

FIGURE 2 MALDI-TOF MS (from reference 2). The target plate is placed into the mass spectrometer. Spots to be analyzed are shot by a laser, desorbing microbial and matrix molecules from the target plate. Charge is transferred from matrix to microbial molecules, and the ionized molecules are accelerated through a positively charged electrostatic field into the mass analyzer, a tube under vacuum. The ions travel toward an ion detector with the smallest analytes traveling fastest, followed by progressively larger analytes. As ions emerge from the mass analyzer, they run into the ion detector, thereby generating a mass spectrum representing the number of ions hitting the detector over time. Although separation is by mass-to-charge ratio, since the charge is typically single for the described application, separation is by molecular weight. Reproduced by permission of Mayo Foundation for Medical Education and Research. All rights reserved. doi:10.1128/9781555817381.ch4.f2

species, but with appropriate library construction, MALDITOF MS should be able to identify most clinicallyrelevant species, with a few caveats. Members of the Mycobacterium tuberculosis complex will likely be identifiable at the complex level only (66), and some species (e.g., Mycobacterium abscessus and Mycobacterium massiliense; Mycobacterium mucogenicum and Mycobacterium phocaicum; and Mycobacterium chimaera and Mycobacterium intracellulare) may not be well differentiated from one another (66). With enhanced databases, Nocardia species (67) may be identified, but as with Mycobacterium species, specific extraction procedures may be required. MALDI-TOF MS can identify yeast (68), outperforming some conventional phenotypic systems, and distinguishing Candida dubliniensis and Candida albicans; Candida rugosa and Candida pararugosa; Candida norvegensis, Candida krusei, and Candida inconspicua; Candida parapsilosis, Candida orthopsilosis, and Candida metapsilosis (69); and (with database supplementation) Cryptococcus neoformans and

Cryptococcus gattii (70–72). Dhiman et al. evaluated the Bruker system for identification of 138 common and 103 unusual yeast isolates and reported 96 and 85% accurate species-level identification, respectively (73). Although older studies used preparatory extraction for yeasts, we and others have used a direct colony testing strategy with formic acid overlay (Fig. 1) (34, 74). Westblade et al. performed a multicenter study assessing the Vitek MS system v2.0 for identification of 852 yeast isolates, including Candida species, Cryptococcus neoformans, and other clinically relevant yeasts, using direct application to a target plate followed by a formic acid overlay; 96.6% were identified to the genus level and 96.1% to the species level (74). MALDI-TOF MS may outperform current systems for esoteric species, such as Candida famata (69). Filamentous fungi exhibit variable phenotypes and protein spectra may vary with growth conditions and with the zone of fungal mycelium analyzed; few are represented in current commercial databases. De Carolis et al. developed a library of Aspergillus species, Fusarium species, and Mucorales using the Biotyper system and identified 97% of 94 isolates to the species level (75). Using the VITEK MS v1 system/ v1.1 database and direct on-plate testing, Iriart et al. identified 82% of 44 Aspergillus isolates (including all isolates with species in the database) (76). With appropriate database building, dermatophytes can be identified (77, 78). Preliminary studies indicate that Pseudallescheria/Scedosporium complex species are identifiable (79). Lau et al. used a special fungal extraction procedure and their own mass spectral database comprising 294 isolates representing 76 genera and 152 species to test 421 mold isolates; they achieved accurate species- and genus-level identifications with 88.9% and 4.3% of isolates, respectively (80). Turnaround time for MALDI-TOF MS is 3 or fewer minutes per isolate. Compared to standard methods, turnaround time for bacterial and fungal identification is shorter by an average of 1.45 days (81), and since only a small amount of organism is required, testing can be performed on single colonies on primary culture plates without subculture. MALDI-TOF MS has a low reagent cost (81), and compared to conventional phenotypic identification and sequencing, reduces costs by 5- and 96-fold, respectively (37). An estimated 87% of isolates may be identified on the first day (compared with 9% with standard techniques), with final identifications several days earlier for biochemically inert, fastidious, or slow-growing organisms (81). DNA sequencing expenses can be avoided for some esoteric organisms, waste disposal decreased, and tests for screening for certain enteric pathogens (e.g., triple sugar iron agar for Salmonella species) and quality control and laboratory technologist training/labor for replaced/retired tests eliminated (82, 83). MALDI-TOF MS has several limitations. Unlike publicly available sequence databases such as GenBank, MALDI-TOF MS databases are proprietary. Although low identification percentages for some organisms may be improved by user addition of mass spectral entries of underrepresented species or strains (to cover intraspecies variability), or even readdition of reference strain spectra to the database, doing so may be beyond the capability of some laboratories. Because of low scores/percentages, repeat testing may be required for ~10% of isolates (81). Growth on some media may be associated with low scores/percentages (84), and small or mucoid colonies may fail identification (40, 85). Refined criteria may be needed to distinguish closely related species and differentiate them from the next best taxon match (86). For certain species of organisms, genus- or

4. Systems for Identification of Bacteria and Fungi n 39 TABLE 4

Evaluations of MALDI-TOF MS for routine bacterial identification (2) Isolates

System No.

Type

Bruker Biotyper

1,013

Bacteria

Bruker Biotyper

468

Bruker Biotyper Vitek MSa Bruker Biotyper

% Identification to level: Period of isolate collection

Country Genus

Species

2 mo

99

97

France

Bacteria

3 mo

97

92

Japan

2,781

Bacteria

1 mo

96

85

Australia

767 986

Bacteria Bacteria

6 wk 3 mo

95 96

87 93

France Belgium

94

93

Vitek MSb

Comparator

Reference

Phoenix, API, Biochemical MicroScan, API, Phoenix VITEK2, API, Biochemical VITEK2 Bruker Biotyper compared to Vitek MS

94 95 96 97 1

a

v1 system/v1.1 database. b Prerelease version of v1.1 database.

species-specific (including lowered) cutoffs may be appropriate (51, 59, 87). Errors may occur, including colony inoculation in erroneous target plate locations, testing impure colonies, smearing between spots, failure to clean target plates, and wrong result entry into laboratory information systems. There is a learning curve to applying ideal colony amounts to target plates (40). Although results are generally reproducible, sources of variability include the mass spectrometer, matrix and solvent composition, technologists, culture conditions, and biological variability; quality control strategies are incompletely developed. Instrument (e.g., laser) and software failure may occur.

GENOTYPIC IDENTIFICATION SYSTEMS DNA Target Sequencing DNA target sequencing may be performed on organisms growing in pure culture. For a comprehensive review, refer to articles and guidelines that specifically address this topic (88, 89). The selection of DNA targets to identify bacteria and fungi relies on the concept that some genes have conserved segments flanking variable regions. Conserved regions of gene targets are locations where PCR and DNA sequencing primers anneal. Variable regions have unique nucleotide sequences, enabling sequence-based identification of a particular genus and species. The gene target most commonly used for bacterial identification is the 16S rRNA gene (16S ribosomal DNA), an ~1,500 bp gene that encodes a portion of the 30S ribosomal subunit. Partial (500 bp) 16S rRNA gene sequencing is commonly used for sequencebased identification of Gram-negative and Gram-positive bacteria, anaerobic bacteria, and mycobacteria (88). For genera with high conservation of the partially sequenced 16S rRNA gene, full-length 16S rRNA gene sequencing or sequencing of an alternative DNA target may be useful. Examples of alternative DNA targets for sequence-based bacterial identification include rpoB (the β-subunit of bacterial RNA polymerase), tuf (elongation factor Tu), gyrA or gyrB (gyrase A or B), sodA (manganese-dependent superoxide dismutase), and heat shock proteins. Although alternative targets may provide better discrimination between species than the 16S rRNA gene, because they are less conserved, genus- or group-specific PCR and/or DNA sequencing primers may be required, and there may be less available sequence data than exists for the 16S rRNA gene.

Potential targets for identification of yeasts and medically relevant molds include the internal transcribed spacer regions ITS1 and ITS2, which are variable regions located between conserved genes encoding 18S, 5.8S, and 28S rRNAs, and the D1-D2 region of 28S rRNA. To identify the microorganism, its DNA sequence is compared to reference sequences found in public (e.g., GenBank) and/or private (e.g., MicroSeq [Applied Biosystems, Calrsbad, CA], SmartGene [Lausanne, Switzerland]) databases. After comparing the query and reference sequences, the number of nucleotide mismatches between the query and reference sequences is used to determine relatedness, and the final result is reported as percent identity. The acceptable percent identity to identify a microorganism to the genus or species level is variable and depends on the DNA target and microorganism. 16S rRNA genes are multicopy targets in most bacteria, and variations in sequence amongst 16S rRNA genes in single organisms can result in difficultly interpreting sequence data. A computer program called RipSeq Mixed (iSentio, Bergen, Norway) can be used to computationally decatenate underlying sequences in such cases. Nucleotide databases must be carefully evaluated for accuracy, quality of sequence data, frequencies of database updates, software, cost, and breadth of nucleotide entries. The Clinical and Laboratory Standards Institute published a comprehensive consensus document for identifying microorganisms to the genus and species levels by DNA target sequencing; this document can serve as a useful guide for laboratorians who wish to pursue or have already implemented DNA target sequencing (88). In the future, wholegenome sequencing will become more routine for clinical laboratories, providing information about the diversity of species, their virulence properties, epidemiology, and antimicrobial resistance mechanisms, and disease causation.

PCR Electrospray Ionization Mass Spectrometry Another technique to characterize bacteria is the application of electrospray ionization-MS to analyze PCR products (PCR/ESI-MS) (90–92). Rather than measuring the mass of bacterial proteins, masses of amplified DNA from broadrange (e.g., rRNA) and specific genes are measured. The masses of amplified DNA are measured with sufficient accuracy to enable unambiguous calculation of the nucleotide compositions of the amplified DNA. Typically, a number of PCR assays are performed and analyzed as a panel. By considering which PCR assays yield amplification products, along with their nucleotide compositions, the identity of

40

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

the microorganism may be defined. This is conceptually similar to broad-range PCR followed by sequencing, except that here MS provides the base composition without knowing the order of the nucleotides. Analyzing multiple target sites on the microbial genome can compensate the lower information content of nucleotide composition over sequence. Simner et al. recently evaluated the PLEX-ID Broad Fungal assay (Abbott Laboratories, Abbott Park, IL) for identification of 91 characterized fungal isolates; 95.6% and 81.3% were identified to the genus and species levels, respectively (93). The advantage of PCR/ESI-MS over sequencing is that it is fast and has a high throughput; however, it is currently quite costly. PCR/ESI-MS is being developed as IRIDICA by Abbott (Abbott Park, IL).

REFERENCES 1. Martiny D, Busson L, Wybo I, El Haj RA, Dediste A, Vandenberg O. 2012. Comparison of the Microflex LT and Vitek MS systems for routine identification of bacteria by matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 50:1313–1325. 2. Patel R. 2013. MALDI-TOF mass spectrometry: transformative proteomics for clinical microbiology. Clin Chem 59:340–342. 3. Willcox WR, Lapage SP, Bascomb S, Curtis MA. 1973. Identification of bacteria by computer: theory and programming. J Gen Microbiol 77:317–330. 4. D’Amato RF, Holmes B, Bottone EJ. 1981. The systems approach to diagnostic microbiology. Crit Rev Microbiol 9:1–44. 5. Edberg S, Konowe L. 1982. A systematic means to conduct a microbiology evaluation, p. 268–299. In Lorian V (ed), Significance of Medical Microbiology in the Care of Patients, 2nd ed. The Williams & Wilkins Co, Baltimore, MD. 6. Sharp S (ed). 2009. Cumitech 31A: Verification and Validation of Procedures in the Clinical Microbiology Laboratory. ASM Press, Washington, DC. 7. Miller JM. 1991. Evaluating biochemical identification systems. J Clin Microbiol 29:1559–1561. 8. Clinical and Laboratory Standards Institute. 2008. User Protocol for Evaluation of Qualitative Test Performance; Approved Guideline, 2nd ed. Clinical and Laboratory Standards Institute, Wayne, PA. 9. Stager CE, Davis JR. 1992. Automated systems for identification of microorganisms. Clin Microbiol Rev 5:302–327. 10. Donay JL, Mathieu D, Fernandes P, Pregermain C, Bruel P, Wargnier A, Casin I, Weill FX, Lagrange PH, Herrmann JL. 2004. Evaluation of the automated Phoenix system for potential routine use in the clinical microbiology laboratory. J Clin Microbiol 42:1542–1546. 11. Linde HJ, Neubauer H, Meyer H, Aleksic S, Lehn N. 1999. Identification of Yersinia species by the Vitek GNI card. J Clin Microbiol 37:211–214. 12. Federal Register. 1992. Clinical Laboratory Improvement Amendments of 1988. Fed Regist 57:7164. http://www.gpo .gov/fdsys/pkg/STATUTE-102/pdf/STATUTE-102Pg2903.pdf. 13. Crowley E, Bird P, Fisher K, Goetz K, Boyle M, Benzinger MJ Jr, Juenger M, Agin J, Goins D, Johnson RL. 2012. Evaluation of the VITEK 2 gram positive (GP) microbial identification test card: collaborative study. J AOAC Int 95:1425–1432. 14. Jin WY, Jang SJ, Lee MJ, Park G, Kim MJ, Kook JK, Kim DM, Moon DS, Park YJ. 2011. Evaluation of VITEK 2, MicroScan, and Phoenix for identification of clinical isolates and reference strains. Diagn Microbiol Infect Dis 70:442–447. 15. Chatzigeorgiou KS, Sergentanis TN, Tsiodras S, Hamodrakas SJ, Bagos PG. 2011. Phoenix 100 versus Vitek 2 in the identification of gram-positive and gram-negative

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26. 27.

28.

29. 30.

31.

32.

33.

bacteria: a comprehensive meta-analysis. J Clin Microbiol 49:3284–3291. Rennie RP, Brosnikoff C, Turnbull L, Reller LB, Mirrett S, Janda W, Ristow K, Krilcich A. 2008. Multicenter evaluation of the Vitek 2 anaerobe and Corynebacterium identification card. J Clin Microbiol 46:2646–2651. Martiny D, Dediste A, Debruyne L, Vlaes L, Haddou NB, Vandamme P, Vandenberg O. 2011. Accuracy of the API Campy system, the Vitek 2 Neisseria-Haemophilus card and matrix-assisted laser desorption ionization timeof-flight mass spectrometry for the identification of Campylobacter and related organisms. Clin Microbiol Infect 17:1001–1006. Siemens Healthcare Diagnostics. 2011. Microscan Dried Gram Negative Procedure Manual, p. 1–20. Siemens Healthcare Diagnostics, West Sacramento, CA. Siemens Healthcare Diagnostics. 2008. Rapid Gram Negative ID Type 3 Procedural and QC Manual, p. 1–3. Siemens Healthcare Diagnostics, West Sacramento, CA. Clinical and Laboratory Standards Institute. 2009. Analysis and Presentation of Cumulative Antimicrobial Susceptibility Test Data; Approved Guideline M39-A3, 3rd ed. Clinical and Laboratory Standards Institute, Wayne, PA. Eigner U, Schmid A, Wild U, Bertsch D, Fahr AM. 2005. Analysis of the comparative workflow and performance characteristics of the VITEK 2 and Phoenix systems. J Clin Microbiol 43:3829–3834. Junkins AD, Arbefeville SS, Howard WJ, Richter SS. 2010. Comparison of BD Phoenix AP Workflow with Vitek 2. J Clin Microbiol 48:1929–1931. Staneck JL, Weckbach LS, Tilton RC, Zabransky RJ, Bayola-Mueller L, O’Hara CM, Miller JM. 1993. Collaborative evaluation of the Radiometer Sensititre AP80 for identification of gram-negative bacilli. J Clin Microbiol 31:1179–1184. Garza-Gonzalez E, Morfin-Otero R, Macedo P, Gonzalez GM, Llaca-Diaz JM, Perez-Gomez R, Rodriguez-Noriega E. 2010. Evaluation of Sensititre plates for identification of clinically relevant coagulase-negative staphylococci. J Clin Microbiol 48:963–965. Morgan MC, Boyette M, Goforth C, Sperry KV, Greene SR. 2009. Comparison of the Biolog OmniLog identification system and 16S ribosomal RNA gene sequencing for accuracy in identification of atypical bacteria of clinical origin. J Microbiol Methods 79:336–343. Biolog. 2012. Biolog Catalogue and Product Guide. Biolog, Hayward, CA. Miller JM, Biddle JW, Quenzer VK, McLaughlin JC. 1993. Evaluation of Biolog for identification of members of the family Micrococcaceae. J Clin Microbiol 31:3170– 3173. Holmes B, Costas M, Ganner M, On SL, Stevens M. 1994. Evaluation of Biolog system for identification of some gram-negative bacteria of clinical importance. J Clin Microbiol 32:1970–1975. Buyer JS. 2002. Identification of bacteria from single colonies by fatty acid analysis. J Microbiol Methods 48:259–265. Van den Velde S, Lagrou K, Desmet K, Wauters G, Verhaegen J. 2006. Species identification of corynebacteria by cellular fatty acid analysis. Diagn Microbiol Infect Dis 54:99–104. Patel R. 2013. Matrix-assisted laser desorption ionizationtime of flight mass spectrometry in clinical microbiology. Clin Infect Dis 57:564–572. Seng P, Drancourt M, Gouriet F, La Scola B, Fournier PE, Rolain JM, Raoult D. 2009. Ongoing revolution in bacteriology: routine identification of bacteria by matrixassisted laser desorption ionization time-of-flight mass spectrometry. Clin Infect Dis 49:543–551. Schulthess B, Brodner K, Bloemberg GV, Zbinden R, Bottger EC, Hombach M. 2013. Identification of grampositive cocci by use of matrix-assisted laser desorption

4. Systems for Identification of Bacteria and Fungi n 41

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

ionization-time of flight mass spectrometry: comparison of different preparation methods and implementation of a practical algorithm for routine diagnostics. J Clin Microbiol 51:1834–1840. Theel ES, Schmitt BH, Hall L, Cunningham SA, Walchak RC, Patel R, Wengenack NL. 2012. Formic acidbased direct, on-plate testing of yeast and Corynebacterium species by Bruker Biotyper matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 50:3093–3095. McElvania Tekippe E, Shuey S, Winkler DW, Butler MA, Burnham CA. 2013. Optimizing identification of clinically relevant gram-positive organisms by use of the Bruker Biotyper matrix-assisted laser desorption ionizationtime of flight mass spectrometry system. J Clin Microbiol 51:1421–1427. Ford BA, Burnham CA. 2013. Optimization of routine identification of clinically relevant gram-negative bacteria by use of matrix-assisted laser desorption ionization-time of flight mass spectrometry and the Bruker Biotyper. J Clin Microbiol 51:1412–1420. Seng P, Abat C, Rolain JM, Colson P, Lagier JC, Gouriet F, Fournier PE, Drancourt M, La Scola B, Raoult D. 2013. Identification of rare pathogenic bacteria in a clinical microbiology laboratory: impact of matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 51:2182–2194. Richter SS, Sercia L, Branda JA, Burnham CA, Bythrow M, Ferraro MJ, Garner OB, Ginocchio CC, Jennemann R, Lewinski MA, Manji R, Mochon AB, Rychert JA, Westblade LF, Procop GW. 2013. Identification of Enterobacteriaceae by matrix-assisted laser desorption/ionization time-of-flight mass spectrometry using the VITEK MS system. Eur J Clin Microbiol Infect Dis 32:1571–1578. Pavlovic M, Konrad R, Iwobi AN, Sing A, Busch U, Huber I. 2012. A dual approach employing MALDI-TOF MS and real-time PCR for fast species identification within the Enterobacter cloacae complex. FEMS Microbiol Lett 328:46–53. Harris P, Winney I, Ashhurst-Smith C, O’Brien M, Graves S. 2012. Comparison of Vitek MS (MALDI-TOF) to standard routine identification methods: an advance but no panacea. Pathology 44:583–585. de Jong E, de Jong AS, Smidts-van den Berg N, Rentenaar RJ. 2013. Differentiation of Raoultella ornithinolytica/planticola and Klebsiella oxytoca clinical isolates by matrix-assisted laser desorption/ionization-time of flight mass spectrometry. Diagn Microbiol Infect Dis 75:431–433. He Y, Li H, Lu X, Stratton CW, Tang YW. 2010. Mass spectrometry biotyper system identifies enteric bacterial pathogens directly from colonies grown on selective stool culture media. J Clin Microbiol 48:3888–3892. Lamy B, Kodjo A, Laurent F. 2011. Identification of Aeromonas isolates by matrix-assisted laser desorption ionization time-of-flight mass spectrometry. Diagn Microbiol Infect Dis 71:1–5. Zangenah S, Ozenci V, Borang S, Bergman P. 2012. Identification of blood and wound isolates of C. canimorsus and C. cynodegmi using VITEK2 and MALDI-TOF. Eur J Clin Microbiol Infect Dis 31:2631–2637. Kuhns M, Zautner AE, Rabsch W, Zimmermann O, Weig M, Bader O, Gross U. 2012. Rapid discrimination of Salmonella enterica serovar Typhi from other serovars by MALDI-TOF mass spectrometry. PLoS One 7:e40004. Lambiase A, Del Pezzo M, Cerbone D, Raia V, Rossano F, Catania MR. 2013. Rapid identification of Burkholderia cepacia complex species recovered from cystic fibrosis patients using matrix-assisted laser desorption ionization time-of-flight mass spectrometry. J Microbiol Methods 92:145–149.

47. Couturier MR, Mehinovic E, Croft AC, Fisher MA. 2011. Identification of HACEK clinical isolates by matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 49:1104–1106. 48. Gaia V, Casati S, Tonolla M. 2011. Rapid identification of Legionella spp. by MALDI-TOF MS based protein mass fingerprinting. Syst Appl Microbiol 34:40–44. 49. Cunningham SA, Patel R. 2013. Importance of using Bruker’s security-relevant library for Biotyper identification of Burkholderia pseudomallei, Brucella species, and Francisella tularensis. J Clin Microbiol 51:1639–1640. 50. Rychert J, Burnham CA, Bythrow M, Garner OB, Ginocchio CC, Jennemann R, Lewinski MA, Manji R, Mochon AB, Procop GW, Richter SS, Sercia L, Westblade LF, Ferraro MJ, Branda JA. 2013. Multicenter evaluation of the Vitek MS matrix-assisted laser desorption ionizationtime of flight mass spectrometry system for identification of gram-positive aerobic bacteria. J Clin Microbiol 51:2225– 2231. 51. Richter C, Hollstein S, Woloszyn J, Kaase M, Gatermann SG, Szabados F. 2012. Evaluation of species-specific score cut-off values for various Staphylococcus species using a MALDI Biotyper-based identification. J Med Microbiol 61:1409–1416. 52. Cherkaoui A, Emonet S, Fernandez J, Schorderet D, Schrenzel J. 2011. Evaluation of matrix-assisted laser desorption ionization-time of flight mass spectrometry for rapid identification of beta-hemolytic streptococci. J Clin Microbiol 49:3004–3005. 53. Senneby E, Nilson B, Petersson AC, Rasmussen M. 2013. Matrix-assisted laser desorption ionization-time of flight mass spectrometry is a sensitive and specific method for identification of aerococci. J Clin Microbiol 51:1303–1304. 54. Davies AP, Reid M, Hadfield SJ, Johnston S, Mikhail J, Harris LG, Jenkinson HF, Berry N, Lewis AM, El-Bouri K, Mack D. 2012. Identification of clinical isolates of alpha-hemolytic streptococci by 16S rRNA gene sequencing, matrix-assisted laser desorption ionization-time of flight mass spectrometry using MALDI Biotyper, and conventional phenotypic methods: a comparison. J Clin Microbiol 50:4087–4090. 55. Alatoom AA, Cunningham SA, Ihde SM, Mandrekar J, Patel R. 2011. Comparison of direct colony method versus extraction method for identification of gram-positive cocci by use of Bruker Biotyper matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 49:2868–2873. 56. Dubois D, Segonds C, Prere MF, Marty N, Oswald E. 2013. Identification of clinical Streptococcus pneumoniae isolates among other alpha and nonhemolytic streptococci by use of the Vitek MS matrix-assisted laser desorption ionization-time of flight mass spectrometry system. J Clin Microbiol 51:1861–1867. 57. Branda JA, Markham RP, Garner CD, Rychert JA, Ferraro MJ. 2013. Performance of the Vitek MS v2.0 system in distinguishing Streptococcus pneumoniae from nonpneumococcal species of the Streptococcus mitis group. J Clin Microbiol 51:3079–3082. 58. Lopez Roa P, Sanchez Carrillo C, Marin M, Romero F, Cercenado E, Bouza E. 2013. Value of matrix-assisted laser desorption ionization-time of flight for routine identification of viridans group streptococci causing bloodstream infections. Clin Microbiol Infect 19:438–444. 59. Alatoom AA, Cazanave CJ, Cunningham SA, Ihde SM, Patel R. 2012. Identification of non-diphtheriae Corynebacterium by use of matrix-assisted laser desorption ionizationtime of flight mass spectrometry. J Clin Microbiol 50:160– 163. 60. Coltella L, Mancinelli L, Onori M, Lucignano B, Menichella D, Sorge R, Raponi M, Mancini R, Russo C. 2013.

42

61.

62.

63.

64.

65.

66.

67.

68.

69.

70.

71.

72.

73.

74.

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS Advancement in the routine identification of anaerobic bacteria by MALDI-TOF mass spectrometry. Eur J Clin Microbiol Infect Dis 32:1183–1192. Barreau M, Pagnier I, La Scola B. 2013. Improving the identification of anaerobes in the clinical microbiology laboratory through MALDI-TOF mass spectrometry. Anaerobe 22:123–125. Jamal WY, Shahin M, Rotimi VO. 2013. Comparison of two matrix-assisted laser desorption/ionization-time of flight (MALDI-TOF) mass spectrometry methods and API 20AN for identification of clinically relevant anaerobic bacteria. J Med Microbiol 62:540–544. Wybo I, Soetens O, De Bel A, Echahidi F, Vancutsem E, Vandoorslaer K, Pierard D. 2012. Species identification of clinical Prevotella isolates by matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 50:1415–1418. Schmitt BH, Cunningham SA, Dailey AL, Gustafson DR, Patel R. 2013. Identification of anaerobic bacteria by Bruker Biotyper matrix-assisted laser desorption ionizationtime of flight mass spectrometry with on-plate formic acid preparation. J Clin Microbiol 51:782–786. Garner O, Mochon A, Branda J, Burnham C-A D, Bythrow M, Ferraro M, Ginocchio C, Jennemann R, Manji R, Procop G, Richter S, Rychert J, Sercia L, Westblade L, M Lewinski. 2014. Multi-centre evaluation of mass spectrometric identification of anaerobic bacteria using the VITEK®MS system. Clin Microbiol Infect 20:335–339. Saleeb PG, Drake SK, Murray PR, Zelazny AM. 2011. Identification of mycobacteria in solid-culture media by matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 49:1790–1794. Verroken A, Janssens M, Berhin C, Bogaerts P, Huang TD, Wauters G, Glupczynski Y. 2010. Evaluation of matrix-assisted laser desorption ionization-time of flight mass spectrometry for identification of Nocardia species. J Clin Microbiol 48:4015–4021. Rosenvinge FS, Dzajic E, Knudsen E, Malig S, Andersen LB, Lovig A, Arendrup MC, Jensen TG, Gahrn-Hansen B, Kemp M. 2013. Performance of matrix-assisted laser desorption-time of flight mass spectrometry for identification of clinical yeast isolates. Mycoses 56:229–235. Castanheira M, Woosley LN, Diekema DJ, Jones RN, Pfaller MA. 2013. Candida guilliermondii and other species of Candida misidentified as Candida famata: assessment by Vitek 2, DNA sequencing analysis, and matrix-assisted laser desorption ionization-time of flight mass spectrometry in two global antifungal surveillance programs. J Clin Microbiol 51:117–124. Firacative C, Trilles L, Meyer W. 2012. MALDI-TOF MS enables the rapid identification of the major molecular types within the Cryptococcus neoformans/C. gattii species complex. PLoS One 7:e37566. Sendid B, Ducoroy P, Francois N, Lucchi G, Spinali S, Vagner O, Damiens S, Bonnin A, Poulain D, Dalle F. 2013. Evaluation of MALDI-TOF mass spectrometry for the identification of medically-important yeasts in the clinical laboratories of Dijon and Lille hospitals. Med Mycol 51:25–32. McTaggart LR, Lei E, Richardson SE, Hoang L, Fothergill A, Zhang SX. 2011. Rapid identification of Cryptococcus neoformans and Cryptococcus gattii by matrixassisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 49:3050–3053. Dhiman N, Hall L, Wohlfiel SL, Buckwalter SP, Wengenack NL. 2011. Performance and cost analysis of matrix-assisted laser desorption ionization-time of flight mass spectrometry for routine identification of yeast. J Clin Microbiol 49:1614–1616. Westblade LF, Jennemann R, Branda JA, Bythrow M, Ferraro MJ, Garner OB, Ginocchio CC, Lewinski MA,

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

Manji R, Mochon AB, Procop GW, Richter SS, Rychert JA, Sercia L, Burnham CA. 2013. Multicenter study evaluating the Vitek MS system for identification of medically important yeasts. J Clin Microbiol 51:2267–2272. De Carolis E, Posteraro B, Lass-Florl C, Vella A, Florio AR, Torelli R, Girmenia C, Colozza C, Tortorano AM, Sanguinetti M, Fadda G. 2012. Species identification of Aspergillus, Fusarium and Mucorales with direct surface analysis by matrix-assisted laser desorption ionization time-offlight mass spectrometry. Clin Microbiol Infect 18:475–484. Iriart X, Lavergne RA, Fillaux J, Valentin A, Magnaval JF, Berry A, Cassaing S. 2012. Routine identification of medical fungi by the new Vitek MS matrix-assisted laser desorption ionization-time of flight system with a new timeeffective strategy. J Clin Microbiol 50:2107–2110. Theel ES, Hall L, Mandrekar J, Wengenack NL. 2011. Dermatophyte identification using matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 49:4067–4071. Nenoff P, Erhard M, Simon JC, Muylowa GK, Herrmann J, Rataj W, Graser Y. 2013. MALDI-TOF mass spectrometry—a rapid method for the identification of dermatophyte species. Med Mycol 51:17–24. Coulibaly O, Marinach-Patrice C, Cassagne C, Piarroux R, Mazier D, Ranque S. 2011. Pseudallescheria/Scedosporium complex species identification by matrix-assisted laser desorption ionization time-of-flight mass spectrometry. Med Mycol 49:621–626. Lau AF, Drake SK, Calhoun LB, Henderson CM, Zelazny AM. 2013. Development of a clinically comprehensive database and a simple procedure for identification of molds from solid media by matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 51:828–834. Tan KE, Ellis BC, Lee R, Stamper PD, Zhang SX, Carroll KC. 2012. Prospective evaluation of a matrix-assisted laser desorption ionization-time of flight mass spectrometry system in a hospital clinical microbiology laboratory for identification of bacteria and yeasts: a bench-by-bench study for assessing the impact on time to identification and costeffectiveness. J Clin Microbiol 50:3301–3308. Gaillot O, Blondiaux N, Loiez C, Wallet F, Lemaitre N, Herwegh S, Courcol RJ. 2011. Cost-effectiveness of switch to matrix-assisted laser desorption ionization-time of flight mass spectrometry for routine bacterial identification. J Clin Microbiol 49:4412. Sparbier K, Weller U, Boogen C, Kostrzewa M. 2012. Rapid detection of Salmonella sp. by means of a combination of selective enrichment broth and MALDI-TOF MS. Eur J Clin Microbiol Infect Dis 31:767–773. Anderson NW, Buchan BW, Riebe KM, Parsons LN, Gnacinski S, Ledeboer NA. 2012. Effects of solid-medium type on routine identification of bacterial isolates by use of matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 50:1008–1013. Xiao D, Zhao F, Lv M, Zhang H, Zhang Y, Huang H, Su P, Zhang Z, Zhang J. 2012. Rapid identification of microorganisms isolated from throat swab specimens of community-acquired pneumonia patients by two MALDITOF MS systems. Diagn Microbiol Infect Dis 73:301–307. De Bel A, Wybo I, Vandoorslaer K, Rosseel P, Lauwers S, Pierard D. 2011. Acceptance criteria for identification results of gram-negative rods by mass spectrometry. J Med Microbiol 60:684–686. Szabados F, Tix H, Anders A, Kaase M, Gatermann SG, Geis G. 2012. Evaluation of species-specific score cutoff values of routinely isolated clinically relevant bacteria using a direct smear preparation for matrix-assisted laser desorption/ionization time-of-flight mass spectrometrybased bacterial identification. Eur J Clin Microbiol Infect Dis 31:1109–1119.

4. Systems for Identification of Bacteria and Fungi n 43 88. Clinical and Laboratory Standards Institute. 2008. Interpretive Criteria for Identification of Bacteria and Fungi by DNA Target Sequencing, vol. MM-18A. Clinical and Laboratory Standards Institute, Wayne, PA. 89. Petti CA. 2007. Detection and identification of microorganisms by gene amplification and sequencing. Clin Infect Dis 44:1108–1114. 90. Baldwin CD, Howe GB, Sampath R, Blyn LB, Matthews H, Harpin V, Hall TA, Drader JJ, Hofstadler SA, Eshoo MW, Rudnick K, Studarus K, Moore D, Abbott S, Janda JM, Whitehouse CA. 2009. Usefulness of multilocus polymerase chain reaction followed by electrospray ionization mass spectrometry to identify a diverse panel of bacterial isolates. Diagn Microbiol Infect Dis 63:403–408. 91. Ecker DJ, Sampath R, Blyn LB, Eshoo MW, Ivy C, Ecker JA, Libby B, Samant V, Sannes-Lowery KA, Melton RE, Russell K, Freed N, Barrozo C, Wu J, Rudnick K, Desai A, Moradi E, Knize DJ, Robbins DW, Hannis JC, Harrell PM, Massire C, Hall TA, Jiang Y, Ranken R, Drader JJ, White N, McNeil JA, Crooke ST, Hofstadler SA. 2005. Rapid identification and strain-typing of respiratory pathogens for epidemic surveillance. Proc Natl Acad Sci USA 102:8012–8017. 92. Ecker DJ, Sampath R, Massire C, Blyn LB, Hall TA, Eshoo MW, Hofstadler SA. 2008. Ibis T5000: a universal

93.

94.

95.

96.

97.

biosensor approach for microbiology. Nat Rev Microbiol 6:553–558. Simner PJ, Uhl JR, Hall L, Weber MM, Walchak RC, Buckwalter S, Wengenack NL. 2013. Broad-range direct detection and identification of fungi by use of the PLEXID PCR-electrospray ionization mass spectrometry (ESIMS) system. J Clin Microbiol 51:1699–1706. Bessède E, Angla-Gre M, Delagarde Y, Sep Hieng S, Ménard A, Mégraud F. 2011. Matrix-assisted laser-desorption/ionization biotyper: experience in the routine of a University hospital. Clin Microbiol Infect 17:533–538. Sogawa K, Watanabe M, Sato K, Segawa S, Ishii C, Miyabe A, Murata S, Saito T, Nomura F. 2011. Use of the MALDI BioTyper system with MALDI-TOF mass spectrometry for rapid identification of microorganisms. Anal Bioanal Chem 400:1905–1911. Neville SA, Lecordier A, Ziochos H, Chater MJ, Gosbell IB, Maley MW, van Hal SJ. 2011. Utility of matrix-assisted laser desorption ionization-time of flight mass spectrometry following introduction for routine laboratory bacterial identification. J Clin Microbiol 49:2980–2984. Dubois D, Grare M, Prere MF, Segonds C, Marty N, Oswald E. 2012. Performances of the Vitek MS matrixassisted laser desorption ionization-time of flight mass spectrometry system for rapid identification of bacteria in routine clinical microbiology. J Clin Microbiol 50:2568–2576.

Automation and Design of the Clinical Microbiology Laboratory CHRISTOPHER D. DOERN AND MARTIN HOLFELDER

5 Clinical microbiology laboratories (CML) are central to the operations of any health care system and take many different forms depending on the type of system they serve. There is no one right way to design a laboratory because appropriate design will be dictated by the space available, staffing requirements, services provided, test volume, and many other factors. Despite institution-to-institution differences in laboratory design, a few elements should be common to all laboratories. First, the laboratory should be a safe environment for employees and visitors. The microbiology laboratory can be a dangerous environment, as the diagnosis of infectious diseases often requires that pathogens be propagated to high concentrations and processed for identification. Laboratory design should be optimized to ensure that the diagnostic process can take place as efficiently and safely as possible. Guidance for safely operating and designing a laboratory can come from many resources. The Centers for Disease Control first published the Biosafety in Microbiological and Biomedical Laboratories in 1984. Although the information provided in this document is advisory in nature, legislation and regulations have in some cases made compliance with this document mandatory (1). At baseline, all CMLs must meet biosafety level 2 criteria and, depending on the services rendered and pathogens encountered, may need to meet biosafety level 3 criteria (1). Second, laboratories should be designed to efficiently handle specimens from initial processing to final result without contaminating the specimen or culture. The efficiency with which a specimen can be processed is an increasing challenge as the centralized laboratory model becomes more common. In these models, a central laboratory provides testing for external institutions that ship specimens to a single location. This model helps to reduce redundancy of resources but at the possible expense of testing efficiency. The time and manner of transport within a hospital and within a system should be considered when designing a laboratory so as to optimize turnaround time (TAT) and reduce contamination. Third, laboratories should be designed so that their configuration can be flexible and accommodate emerging technologies and changing demands. The goal of this chapter is to provide information regarding the proper design of a laboratory, bearing in mind that each institution requires something different from its labora-

tory. Microbiology is rapidly changing and so too are the spatial and geographical demands placed on facilities. The chapter is presented in two sections. The first will focus on laboratory design and workflow. This section will focus on some key elements of laboratory design, including the geography of a laboratory (i.e., where testing is performed), preanalytical considerations, staffing strategies, workflow, process improvement, and clinical impact. The second will address a new and important subject for clinical microbiologists, laboratory automation. Some historical perspective will be provided along with a discussion of currently available options for laboratory automation and considerations for implementation.

GEOGRAPHY OF THE LABORATORY Almost every clinical service is a customer of the laboratory. It is therefore critical that the laboratory be well connected, both electronically and physically, to the facilities for which it provides testing. A wide variety of laboratory information systems (LISs) and electronic medical records can be found among institutions. These systems are now a vital component of laboratory testing and will likely become even more important as digital microbiology and total laboratory automation (TLA) become more common. A review of these numerous systems is beyond the scope of this chapter. The Association for Pathology Informatics provides a toolkit which can be used to assess currently available LISs (http:// www.pathologyinformatics.org/toolkit). In any health care setting, but especially teaching hospitals, convenient access to the laboratory is an important factor that encourages physicians and trainees to interact with the hospital directly. It is difficult to quantify the clinical benefit of this interaction, but most microbiologists and infectious disease practitioners agree that being able to readily access the laboratory for personal consultation is of great benefit. Clearly, there can be real advantages to locating a laboratory within the hospital it serves. However, there can be significant benefits to the centralized model as well. The primary advantage is one of scale. Centralizing a laboratory service, while cumbersome from the perspective of specimen transport, simplifies spatial, technological, and staffing needs. By centralizing services, laboratories can focus all of their resources in a single area, thus minimizing the

doi:10.1128/9781555817381.ch5

44

5. Laboratory Design and Workflow n 45

significant costs of maintaining redundant laboratory infrastructure in external laboratories. Furthermore, in an age where more laboratorians are retiring than entering the work force, highly skilled expertise can be hard to find (2). Where expertise is sparse, centralization allows for many laboratories to take advantage of the skills of the very few. For example, in the United States, parasitic disease is rare. The diagnostic methodology of choice for most parasitic gastrointestinal disease (other than Cryptosporidium and Giardia spp.) is a microscopic ovum and parasite exam. This testing is time-consuming and difficult to maintain proficiency due to low prevalence. Many laboratories have realized they can no longer justify performing this testing in-house because they simply don’t have the expertise. As a result, it is estimated that nearly 60% of laboratories in the United States currently send their ovum and parasite exams to a reference laboratory (3).

LOCATION OF TESTING Traditional models of laboratory structure, where the analyte, rather than the test method, dictated the location of a given test, are now being challenged. There are many examples of infectious disease testing that now occur in the clinical chemistry lab or point-of-care tests that are performed in the Emergency Department rather than in the CML. As diagnostic methods are simplified, it is possible that home testing will become more common.

INFECTIOUS DISEASE TESTING IN THE CHEMISTRY AND CORE LABORATORIES Technology continues to advance and simplify the performance of testing. Many tests that had previously existed in one section of the lab can now be moved to the location of greatest convenience. A good example of this phenomenon is the simplification of molecular testing. Instruments like the GeneXpert (Cepheid, Sunnydale, CA) and the FilmArray (Biofire Diagnostics, Inc., Salt Lake City, UT) have reduced molecular biology to simple specimen manipulation. It is no longer necessary to have skilled molecular biologists performing these tests. This allows institutions to consider relocating testing to a section of the laboratory that may not have previously been an option. Some microbiology laboratories are not staffed 24 h/day, 7 days/week (24/7), and in those scenarios, these “walk-away” molecular tests could be moved to a section of the laboratory that could handle them around the clock if necessary. Deciding on the location of point-of-care testing (POCT) can be a contentious and controversial process. Traditionally, microbiologists have resisted requests to allow nonlaboratorians (such as Emergency Department providers) to perform POCT on the grounds that testing personnel will not adhere to the rigorous standards of protocol and quality control (QC) utilized by trained laboratory staff. In addition, it is sometimes the case that nonlaboratory departments do not fully understand the complexities of performing even a technically uncomplicated test. Tasks such as proper ordering, resulting, competency and proficiency testing, and inventory management must be considered when determining whether nonlaboratory departments should perform POCT. In cases where this is allowed, a rigorous system to monitor the quality of testing should be put in place. In addition to these practical matters, regulatory considerations may also dictate who can perform tests and where testing can be performed.

One advance that has greatly improved the ability to perform high-quality POCT outside the laboratory is instruments that autoverify results. In some cases, these instruments can be programmed for QC lockout, automated result interpretation, and autoverification of results. These instruments aid busy individuals in reading results at the appropriate time and automatically sending the information to the information management system. Lastly, the QC lockout is a function that forces proper QC testing by locking the instrument until QC is performed and passed.

PREANALYTICAL MICROBIOLOGY A critical aspect of laboratory operations is the interaction with the client. Interactions include ordering, specimen tracking, and result reporting, all of which should be delivered electronically. The most obvious interaction with a laboratory involves specimen transport. There are three main forms of specimen transport utilized: manual, pneumatic tube, and robotic. Hospitals with internal laboratories commonly rely on a combination of all three transport modalities, although for most specimens, pneumatic tubes are often preferred. Transportation is a much bigger challenge for the centralized laboratory model. This applies both to commercial reference laboratories and to hospital systems which serve a network of external clients. Efficient transport of specimens is imperative to providing reliable and accurate results. With labile pathogens present in specimens that constitute less-than-ideal environments for maintaining organism viability, minimizing specimen transport time is critical. Although there is surprisingly little literature addressing the importance of specimen transport time, Table 2 in chapter 18 of this Manual provides guidance on the matter. Table 2 recommends that specimens such as cerebrospinal fluid, blood culture bottles, abscess material, and body fluids all be transported (collection to processing) in less than 2 h (4). In scenarios where specimens must be transported to distant laboratories, whether across town or across the country, it would seem unlikely that these recommendations could be met. In light of this fact, some laboratory systems have elected to inoculate and preincubate select, high-priority cultures (such as cerebrospinal fluid) so they can be transported at regular intervals rather than on a STAT basis. Organism viability is one reason that minimizing transport time is critical. A secondary reason is simply that results can be provided faster when specimens are transported efficiently.

STAFFING MODELS The most important resources in a CML are the personnel that perform testing. It is well documented that there is a shortage of skilled medical technologists entering the field. It is estimated that 39% of United States laboratories have budgeted openings, with 57% of those openings being for medical technologists (MT) or clinical laboratory scientists and 14% for medical laboratory technicians (MLT) (2). It is also estimated that laboratories must hire approximately 12,000 new employees annually to keep up with test volume growth. However, training programs are disappearing and only about 5,000 medical technologists enter the field each year. As a result, institutions utilize a number of different staffing models to optimize the effectiveness of their workforce. Most laboratories break down their workday into three 8-h shifts which generally include a daytime, early evening, and night shift. Another approach is the “7 on, 7 off” or

46

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

“8 on, 6 off” schedule in which employees work 7 or 8 straight days of 10-h shifts and then get 7 or 6 days off, respectively. It is not uncommon for laboratories to use a combination of 8- and 10-h shifts to cover the workload. In many laboratories, the majority of staff are deployed to the first shift where most of the high-complexity testing is conducted. Second and third shift staff in this model may help to complete daytime work but will in many cases be responsible for performing only Gram stain interpretation and STAT laboratory functions. An alternative approach to the traditional model is 24/7 staffing. In these laboratories, staffing is evenly distributed throughout the day and cultures are read continuously. The idea for this model was born out of lean/Six Sigma analyses which observed that some cultures were ready to be read during evening and night shifts. It followed that the TAT was unnecessarily delayed when these cultures weren’t attended to until the following morning. In the 24-h staffing model, cultures are first read approximately 16 to 20 h after inoculation. Benches can then be subdivided by time of reading rather than by patient last name or by specimen source. There is a paucity of peer-reviewed literature addressing the effectiveness of this approach. However, at least one lab has shared its experience with 24-h culture reads and shown that TATs are improved by almost 24 h for positive cultures (5). Some other benefits from this experience appear to be that work is more evenly distributed throughout the day. As a result, the morning shift staff do not get overwhelmed by cultures that have accumulated overnight. Lastly, there may also be some financial benefit in both staffing and supply expenses (J. Campos, personal communication). There are some significant challenges to implementing 24/7 culture reading. It may be difficult to find experienced technologists to work on evening and night shifts. Employee turnover is typically higher on these shifts and may make it difficult to maintain trained personnel. Likewise, these shifts are generally unsupervised and without medical direction. Laboratories will want to ensure that they have the same resources for troubleshooting and guidance as are available during day shifts. Another consideration is whether the laboratory will be over-functioning by moving to 24/7 culture reading. Will results released during off-shifts be acted upon? Lastly, change management when moving to 24/7 microbiology will pose a significant challenge. Laboratories may experience resistance when moving from traditional to 24/7 microbiology.

STAFF TRAINING LEVEL Laboratory staff can have various levels of education and experience which dictate the tasks they can perform. Becoming an MT (also referred to as a clinical laboratory scientist, biomedical scientist, or medical laboratory scientist) generally requires a bachelor’s degree in medical technology or in a science such as biology or chemistry. MT programs are usually offered through a university and involve hospital setting internships, and some states may require that MTs pass an exam to be licensed. MTs are qualified to perform complex testing and analyses, including direct specimen microscopy, culture interpretation, organism identification, and susceptibility testing, to perform high-complexity molecular testing, and to supervise others. MTs are expected to understand the underlying scientific principles of laboratory testing as well as the causes of disease and the importance of test selection (6).

An MLT is defined as someone who has a working comprehension of technical and procedural aspects of laboratory tests. The technician correlates tests with disease processes, understands basic physiology, and recognizes abnormal test results (6). The tasks that can be performed by MLTs will vary by institution but generally include quality assurance monitoring, computer applications, and instrumentation troubleshooting and require an understanding of specimen collection and processing. The American Society of Clinical Pathologists states that a technician may make technical decisions related to testing but should be supervised by a technologist, supervisor, or laboratory director. MLTs usually have an associate degree from a community or junior college or a vocational school. MLTs may become MTs through additional education and experience. Medical laboratory assistants are individuals who receive on-the-job training and may have specialized education, but that is not required. The primary task of the medical laboratory assistant is specimen processing and using preanalytical systems. These professionals must have in-depth knowledge of specimen acceptability.

WORKFLOW Batch Versus Immediate Testing The organization of workflow and the selection and extent of batching procedures depend on several factors. Specimens that have direct diagnostic or therapeutic consequences can be batched only to a limited extent. When analytical process automation is involved to a high degree, specimens can be combined in larger series. The effects on TAT must be determined and accordingly taken into consideration. It should also be determined when the findings should be made available to the clinician. When the above conditions are met, the ideal size of a batch can be determined on the basis of the number of specimens, specimen type, time allotted for transport, distribution of specimens after arrival at the laboratory, desired tests, and selected laboratory equipment. The effects of an order entry system or the various options of digital transmission of the findings must also be taken into account. The use of matrix-assisted laser desorption–ionization time of flight (mass spectrometry) (MALDI-TOF [MS]) and molecular methods for organism identification can shorten the TAT (7). Identification using MALDI-TOF (MS) is possible with only a single colony from the primary culture, and fewer subcultures are needed to obtain a pure culture. It thus becomes possible to identify bacteria 1 to 2 workdays earlier than with current phenotypic methods (8, 9). The TAT can be further shortened by changing the reading intervals for the culture plates. Instead of reading the plates only at a fixed time, traditionally in the morning, they could also be read at regular intervals in the afternoon and/or evening. This is particularly true for specimens arriving at the laboratory in the afternoon or evening. Batching the specimens according to the above reading modalities and deploying staff as described above make it possible to maximize laboratory efficiency. Introduction of digital image processing into diagnostic microbiology offers new opportunities for organizing workflows and shortening TAT. Reading plates digitally results in earlier recognition of colony growth. Consequently, subsequent identification and susceptibility testing can be accelerated. The pictures of the plates are taken at individually defined intervals and times for further processing. The computer can be supportive in preselection of culture-negative

5. Laboratory Design and Workflow n 47

plates. The plates showing growth are sent through further diagnostic procedures. Imaging technology can detect even the smallest of colonies, which are difficult for the human eye to differentiate. Chromogenic media for diagnosing multidrug-resistant organisms such as methicillin-resistant Staphylococcus aureus, vancomycin-resistant enterococci, and multidrug-resistant Escherichia coli, Acinetobacter spp., or Salmonella spp. are particularly suitable for imaging technology, since results of the cultures can be documented earlier. Adapting work hours to conform to the above procedures should be considered. Incubation and digital image processing can also be applied to manually inoculated plates. The benefits of automation can be put to full use by batching the specimens for the process described. The rapid development of easy-to-use molecular diagnostic test systems for detecting pathogens like methicillinresistant S. aureus, Clostridium difficile, enterovirus, and respiratory viruses has led to progressively smaller batch sizes. Batching specimens for such tests would thwart the advantages of these systems (e.g., time savings and easy handling).

Role of Process Improvement Economic pressure and new approaches in the domain of laboratory automation, as described below, open up opportunities for reconsideration and possible revision of the entire laboratory process. Potential areas for improvement range from analyzing and developing solutions for the current laboratory situation, concerning workflow, staff utilization, and certain selected methods, up to introducing entirely new laboratory concepts and equipment. Changes in these areas can lead to increased capacity and flexibility within the laboratory and enable it to deal with unforeseen or planned increased numbers of specimens for processing. Another reason for evaluating the entire laboratory organization is to compensate for a dwindling laboratory work force. For instance, in Germany, the number of graduates from schools for medical technologists decreased from 2,273 between 1994 and 1995 to 1,471 between 2009 and 2010 (10). The following steps should be taken to achieve sustained optimization. First, a list of laboratory processes should be compiled. Then an implementation plan should be drawn up. The plan should describe the individual steps: the necessary resources, organization, productivity, target TAT, and ways to deal with additional workload. Sustainability of the new process is achieved by regularly and closely tracking the implemented changes and by monitoring them to determine whether or not objectives have been achieved. The entire process should be preceded by a comprehensive informatory phase: visits to other laboratories and exchanges with colleagues. One can also seek the advice of external consulting firms. Manufacturers of medical equipment for diagnostic purposes also offer concepts. These firms provide a baseline evaluation and analysis of the laboratory’s equipment in its entirety, including the work organization and the work processes, the stock of devices, and the computer systems and their usage. The proposed solutions generally lead to a greater degree of automation, which, along with potentially recommended middle software, provide better coordination of the devices and processes. Various methods for process analysis and optimization are being used in this context. The methods have been adopted from the automobile and shipping industries and are based on presentations from lean management and Six Sigma. The lean management method originates from the automobile industry and was first used by Toyota. The aim was to identify all of the characteristics that enhance the value

of the company and increase customer satisfaction. At the same time, superfluous activities and processes were to be discontinued. Lean management as it pertains to health care can be described in four components: (i) methods for analyzing processes, and thereby identifying and analyzing problems; (ii) methods for designing processes more effectively and efficiently; (iii) methods for better detecting errors, implementing solutions, and preventing damaging effects; and (iv) methods for managing these changes and problems and for finding solutions using a scientific approach (11). A practical example is the implementation of lean management using the concept of the five Ss: exclusively, very necessary materials are allowed in the work area (sorting), the entire laboratory area is straightened up and each piece of working equipment is assigned its own designated place (straightening), the area is cleaned systematically on a regular basis (sweeping), the processes in the laboratory are standardized (standardizing), and sustainability is achieved through regular evaluation and analysis of these processes (sustaining). The aim of the Six Sigma method is to reduce process variance and simultaneously reduce errors and deviations in the analysis to a minimum. This is achieved through implementation of control mechanisms that link workers, work processes, quality requirements, responsibilities, and costs. The method was developed in the 1970s in Japan for the shipbuilding industry and is used today in many branches of industry worldwide. Six Sigma is based on statistical methods for quality improvement aimed to increase organization success and customer satisfaction. The most frequently used Six Sigma method is the so-called DMAIC cycle (define, measure, analyze, improve, and control). Using DMAIC, processes become measured variables, which can be improved in a sustainable manner. The sigma value describes how often an error is expected to occur. The best sigma value in use is 6; this signifies that fewer than 3.4 defects or errors per million opportunities (DPMO: defects per million opportunities) are expected to occur. A sigma value of 1 for laboratory processes would signify that 691,462 defects are expected to occur per million analyses. A laboratory implementing Six Sigma was able to increase its sigma value from 3.9 (7,210 DPMO) to 4.5 (1,387 DPMO) (12). The methods described above are used in various combinations to improve the laboratory organization as part of the optimization process and the implementation of highgrade automation. There are a number of publications on this subject that describe how this process can be applied successfully in many sectors of health care (11, 13–19). Rutledge et al. demonstrated in a clinical laboratory a decreased walk pattern for technologists by 70% and a mean TAT reduction of more than 50% for creatinine, complete blood count, urine analysis, and ionized calcium after implementing lean management and Six Sigma (13). Persoon et al. used the lean production system from Toyota to improve the preanalytic processes in a clinical chemistry laboratory (20). The median preanalytic processing time was reduced from 29 to 19 min. Overall, these process improvement systems promise to increase staff efficiency and reduce costs and errors with the ultimate goal of improving patient outcomes.

LABORATORY AUTOMATION: HISTORICAL PERSPECTIVES The microbiology laboratory has changed very little over the past 30 years with respect to laboratory automation.

48 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

The field, though, is now poised to undergo a shift to automate what has traditionally been a very manual discipline. Despite the fact that clinical chemistry laboratories adopted TLA over 20 years ago, the concept has been slow to be accepted in microbiology (21). A number of factors have contributed to the delay in automating clinical microbiology laboratories. First, the complexity of the microbiological specimen has hindered the development of TLA solutions. The clinical chemistry specimen is of relatively uniform volume and consistency and, perhaps most importantly, is collected in standardized containers. These factors are conducive to automation and are in sharp contrast to the specimen received in CMLs. The CML must be able to process and analyze an enormous variety of specimen types which are collected in highly variable containers. Second, the nature of diagnosing an infectious disease is complicated by nonspecificity of clinical manifestations. The diagnostic process must therefore include testing for a large number of pathogens, and automated platforms must be able to accommodate this reality. Artificial intelligence programs may one day be able to replicate the complex interpretations required by microbiology technologists. Currently, though, culture interpretation requires human judgment and cannot be automated. Third, the cost of automation is a significant barrier to entry into the CML. Relative to chemistry testing, microbiology specimen volumes are much smaller, thus reducing the need for automation and making for less appealing returns on investment. However, the highvolume, centralized laboratory model is becoming common and may be an attractive setting for automation. How these systems will fit into the smaller laboratory is yet to be determined.

CURRENT SYSTEMS FOR MICROBIOLOGY LAB AUTOMATION Organism Identification and Susceptibility Testing Microbiologists have a number of different options for performing organism identification and susceptibility testing. Susceptibility testing systems are reviewed in chapter 72 of this Manual. MALDI-TOF (MS) is a technology capable of providing identifications based on assessment of protein profiles and database comparison. In actuality, these systems are semiautomated, although they currently require significant manual manipulation prior to analysis. The two major platforms for MALDI-TOF (MS) organism identification are the Vitek MS (bioMérieux) and the Biotyper (Bruker Daltonic, Billerica, MA). These are discussed in more detail in chapter 4.

Molecular Automation Traditional molecular biology is a tedious and labor-intensive process requiring experience and great care to prevent contamination. The future of molecular biology is going to look very different as manual molecular processes are being automated and consolidated into single systems. These systems are commonly referred to as being “sample-to-answer” or “walk-away.” In addition to fully automated systems, a number of manufacturers offer automated nucleic acid extraction platforms. Discussing each product individually is outside the scope of this chapter. Automated molecular testing platforms are reviewed in chapter 6 of this Manual.

Automated Specimen Processing Automated inoculation of clinical specimen has been the center of research and development for several years (22). Just recently, these devices became available on the market. Automation of microbiological specimen processing promises to improve the quality of the streaking process, avoid cross-contamination, alleviate ergonomic issues, and reduce processing time and costs. It is expected that the automated streaking process will be reproducible and reliably yield isolated colonies. This will reduce the number of subcultures necessary for identification and susceptibility testing. Available devices have been scientifically evaluated in only a few cases (23–26). The systems usually process swabs in liquid transport media such as E-Swab (Copan Diagnostics, Murrieta, CA) or Sigma swab (Medical Wire, Corsham, United Kingdom). These swab systems consist either of an open-pore polyurethane foam tip and a modified Amies medium (Sigma swab) or a nylon flocked swab with Amies medium (E-Swab). These systems lead to a significantly improved transition of microorganisms from the swab to the transport medium and allow better evaluation of the microorganisms from the Gram stain (20, 25). Swabs in solid or half-solid transport media can only be processed in semiautomatic devices. The devices available are divided according to different inoculation techniques using the loop, comb applicator, or bead technique. In Table 1, the automated inoculation systems are compared regarding specimen type processed, inoculation technique, capacity (inoculated plates/hour), and additionally required disposables.

Innova Innova (Becton Dickinson, Sparks, MD) has five specimen drawers that can accept a total of 200 specimens. A specimen drawer can only accept one type of specimen at a time. The specimens are decapped and recapped automatically and agitated. Up to 270 whole plates as well as biplates of up to six different types can be loaded simultaneously into the device. The streaking pattern can be defined according to the material and selected from a variety of streaking options. The inoculating loop is thermally sterilized. Each loop can be used for up to 15,000 inoculations. For inoculation volumes of 200 µl, a pipette is available. The inoculated plates are sorted according to the type of media into five different groups. The device is a closed system, containing air that is cleaned by a HEPA filter system.

InoqulA FA/MI InoqulA (BD-Kiestra) FA/MI (full automation/manual interaction) can process liquid media in FA mode, and swabs or other types of specimen in MI mode. A barcode is attached to the side of the plates. In MI mode, the swab or the material to be inoculated is placed manually on the agar plate and streaking is performed with magnetic beads as in FA mode. Slides are prepared in MI mode. Processing can be carried out in MI mode or in FA mode but not in both simultaneously. In FA mode, the specimens are automatically agitated as well as decapped and recapped. A magnetic rolling bead (Fig. 1) is used for streaking, and a maximum of five whole plates or biplates can be inoculated at one time. The streaking pattern can be either selected from a variety of patterns or defined by the customer for each material. The inoculated plates can be presorted into four different cassettes for incubation. The system is equipped with a HEPA filter system.

5. Laboratory Design and Workflow n 49 TABLE 1

Automated inoculation systems

Instrument (manufacturer)

Innova (BD-Diagnostics, Sparks, MD) InoqulA FA, MI (BD-Kiestra B.V., Drachten, The Netherlands) PREVI Isola, (bioMérieux, Marcy l’Etoile, France) WASP (Copan Italia Spa, Brescia, Italy)

Inoculation technique

Specimen type

Capacity (no. of inoculated plates according to manufacturer)

Additionally required disposables

Liquid-based specimen

Loop (1, 10, 30 µl)

180a

Liquid-based specimen (FA); swab specimen (MI)

Bead

400a

None; for volumes >200 µl, pipette Pipette, bead

Liquid-based specimen

Comb applicator

180b

Pipette, comb

Liquid-based specimen

Loop (1, 10, 30 µl)

—a

None

a

Capacity depends on the number of inoculated plates or biplates per specimen and chosen streaking pattern. —, capacity depends on the number of inoculated plates or biplates per specimen.

b

PREVI Isola PREVI Isola (bioMérieux, Marcy l’Etoile, France) has five racks designed for various types of specimen. A rack can accept only one type of specimen container at a time. However, the device only processes liquid media. Whole plates and biplates can be loaded into five input cassettes, each of which can accept 30 plates. The specimens must be decapped and recapped manually. The type of streaking pattern is predefined by the comb applicator (Fig. 2). The inoculated plates are stored in three output cassettes (30 plates/cassette). A HEPA filter system is provided.

WASP The WASP (walk-away specimen processor; Copan) processes various specimen types; the specimens are automatically decapped and recapped as well as agitated or centrifuged. Between 342 and 370 whole plates and biplates can be stored in up to nine-plate silos; only one plate type should be used per silo. The streaking pattern can be selected from a variety of options. The inoculating loop is thermally sterilized. The inoculum in the loop is documented per photo. An agar plate can be inoculated one-half each with two different specimens. The inoculated media are sorted according to the plate type. The plates are labeled on the side or on the base. Gram slide preparation, inoculation of enrichment broths, and an antibiotic disk dispenser for susceptibility testing are available. Cultured plates can be reloaded in the WASP for

FIGURE 1 InoqulA magnetic bead. doi:10.1128/9781555817381.ch5.f1

disc diffusion susceptibility testing. The WASP is equipped with a HEPA filter system.

Total Laboratory Automation TLA in microbiology aims to improve quality, reduce time to result, better manage an increasing number of specimens, compensate for reduction in skilled staff, and be more economically effective (27–30). Three companies provide different solutions for TLA in microbiology: BD-Kiestra, bioMérieux, and Copan. The field is evolving quickly. More methods and devices are expected to be automated in the near future. These include automated colony picking for MALDI-TOF MS identification and preparation of dilutions for susceptibility testing.

BD-Kiestra TLA Concept The BD-Kiestra TLA Concept is a conveyer-connected system (Fig. 3). This system comprises the following work steps: inoculation of liquid specimen as well as swabs and other nonliquid specimens using InoqulA, incubation in aerobic and CO2 atmospheres, and digital imaging. Each incubator

FIGURE 2 PREVI Isola-inoculated agar plate. doi:10.1128/9781555817381.ch5.f2

50

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

FIGURE 3 BD-Kiestra TLA with barcoding, inoculation (InoqulA) (depth, 93.5 cm; width, 417.2 cm; weight, 770 kg), and incubation (ReadA) (depth, 164.0 cm; width, 87 cm; weight, 430– 730 kg). doi:10.1128/9781555817381.ch5.f3

has a capacity of 1,152 plates. The plates are stored individually in the incubator. Automated colony picking for identification with MALDI-TOF (MS), automated susceptibility tests, and expansion with molecular diagnostics equipment (BD-Max) are in the planning stage. Actual concepts are individualized with respect to the size and capacity of the installation. In this way, the laboratory can gradually approach full automation and remain open for future developments.

1,000 plates with single-plate location. These instruments can be connected by a conveyor system. At present, measures are being taken to automate the colony picking from agar plates for identification using MALDI-TOF (MS) and susceptibility testing. Additional instruments can be connected to the system using the middle software Myla. These include blood culture systems, Gram-staining devices, Vitek 2, and others.

WASP Lab bioMérieux Concept FMLA The bioMérieux Concept FMLA (full microbiology lab automation) (Fig. 4) is a modular design. The instruments are connected and regulated by the middle software Myla. This system automates the following processes: inoculation of fluid specimen on agar plates using PREVI Isola, incubation in aerobic and CO2 atmospheres, and digital imaging in a smart incubator system. An incubator has a capacity of

The WASP Lab (Copan) (Fig. 5) is a modular construction and connects the individual machines through a conveyerconnected system. The system processes smears, sputum, stool, and liquid samples. It includes specimen processing with the WASP, incubation in aerobic and CO2 atmospheres, and digital imaging. An incubator has a capacity of 800 to 1,760 plates with single-plate location. Preparation is being made to incorporate identification (using MALDI-

FIGURE 4 FMLA, bioMérieux, smart incubation unit (depth, 145.7 cm; width, 128.0 cm; weight, 850 to 950 kg); PREVI Isola (depth, 90.7 cm; width, 144.5 cm; weight, 298 kg). doi:10.1128/9781555817381.ch5.f4

5. Laboratory Design and Workflow n 51

FIGURE 5 WASPLab, Copan: WASP (depth, 119.1 cm; width, 194.9 cm; weight, 700 kg); double incubator (depth, 83.0 cm; width, 174.0 cm; weight, 700 kg); imaging system (depth, 121.3 cm; width, 37.5 cm; weight, 250 kg). doi:10.1128/9781555817381.ch5.f5

TOF [MS]) and automated susceptibility testing into the WASP Lab system.

Digital Imaging Digital imaging is one of the central components in TLA. The methods will be described collectively, since all of the manufacturers adhere to similar principles of image assessment. Digital imaging software is designed to simulate and improve the visual assessment of organism growth. Each system (BD-Kiestra, bioMérieux, and Copan) can take pictures of plates with several exposures and at various angles. With the incorporation of digital imaging and the automation of MALDI-TOF (MS) identifications, the percentage of plates needing manual processing can be greatly reduced. In each system, colonies of bacteria can be labeled for further processing on a screen. Digital processing facilitates early detection of organism growth and shortens the time of identification. Decision making can be automated for certain criteria, e.g., growth or no growth. In addition, all samples from one patient, e.g., urine, sputum, and multidrug-resistant organism screening, can be evaluated simultaneously with computer-assisted processing. Lastly, images can be archived and later used for training or for QC programs.

Limitations of the Systems The strength of the automatic systems described lies in the processing of standardized and uncomplicated specimens. Those specimens needing special processing methods, such as organ or tissue biopsies, are not easily accommodated by such systems. However, the possibilities for automatic processing can be expanded considerably through the use of liquid-based specimen transport systems. This effectively converts a very high volume specimen, the swab, into a

liquid specimen that can be easily managed with an automated specimen processor (25, 27, 31, 32). A high level of flexibility and system compatibility is especially necessary for small laboratories. The modular nature of the automation systems described enables the laboratories to achieve various levels of automation. The combination of modular automation and process improvement, as explained, promises scalability of the current automation systems for laboratories independent of their size. For this reason, the costs of automation for small laboratories may not outweigh the benefits.

Criteria for Evaluation and Selection of an Automation System Prior to choosing an automation system, the daily routine in the laboratory should be evaluated. Data regarding specimen volume, arrival time of specimen, and workflow must be determined to negotiate the future lab design with the manufacturers being considered. If possible, it is recommended that institutions conduct a thorough assessment of the systems installed in other institutions to determine whether or not each system is capable of fulfilling a laboratory’s needs. All types of laboratory personnel should be involved in the project to incorporate their suggestions and ideas as well as to increase general support for the necessary changes. There are only a few publications addressing this issue (27, 29, 30, 33). Factors to consider when selecting an automated system are outlined below.

Productivity The productivity of the automation systems depends upon the number of specimens processed and the TAT for the

52

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

entire analysis process. Productivity is affected by a number of factors. Specimen type and the number of plates per sample must be considered. Respiratory material, urine, wound swabs, genitourinary tract specimens, and screening swabs for multidrug-resistant organisms are processed with different types of media (34). The productivity is influenced by the choice of a streaking pattern, the number of pictures taken by digital imaging, and whether a whole plate or a biplate is inoculated.

the consumable supplies, e.g., pipettes, combs, or beads. It might be necessary to purchase agar plates and broths from different manufacturers. Changes in workflow can lead to alterations in the staffing requirements on different shifts and may necessitate installing a night shift if one does not currently exist. This must be taken into account when budgeting expenses. Service and repair charges, as well as the costs of the interfaces, must also be included. Possible structural changes in the laboratory should also be represented in the financial plan.

Reliability, Stability, and Durability Indicators showing possible malfunction of a system should be used. Contingency plans for possible failures of the system should be planned in advance. Allowance for a certain number of inspections and service procedures should be considered, allotting time for their duration. Staff to service the system should be available on weekends and during holidays.

Technical Aspects In the beginning, do a survey of the buildings and determine the infrastructure of the rooms: calculate the required space and the weight of the machines; determine the power supply and, if necessary, the compressed-air outlets. At present, the dimensions of the devices and systems are continually decreasing in size. Future developments must also be taken into account. The possible integration of more devices, e.g., PCR equipment, should also be considered.

Software Applications The instruments should be connected with a bidirectional interface to the LIS. It might be necessary to use middleware. Coordinating the middleware software and LIS is essential. If required, instruments from other manufacturers should be integrated.

Safety and Hygiene The automated specimen processors should be closed and an appropriate air filter system, e.g., HEPA filter, applied. The risk of contamination must also be taken into account, and protocols for cleaning and disinfecting the incubators and conveyors should be established. Occupational health and safety regulations should be taken into account. There are no current regulations or guidelines addressing the issue of safety in laboratory automation. Of significant concern is the matter of containing exposures generated through errors in automatic incubation.

Quality Control and Scientific Aspects The entire system should be integrated into the quality management program. The individual work steps must be monitored in compliance with the laboratory regulations: from reading the barcode on the samples and plates to inoculating and transporting the plates, digital image processing, and waste disposal. Inoculators should ensure that particles and air bubbles in the specimen are detected to avoid false-negative inoculations. Measures against transposition and cross-contamination of the samples should be ensured. Only one study evaluating cross-contamination by automated systems has been published (24). In this study, sterile and E. coli-inoculated Vacutainer tubes as well as E-Swab tubes were alternately loaded on the WASP. No colonies were observed from the sterile specimen.

Costs Laboratories must take into consideration not only the amount invested in the purchase of the devices, but also

The Final Decision The choice of the type and extent of lab automation is, of course, dependent upon the individual circumstances and the financial resources available. It has been a long time since diagnostic medical microbiology has experienced such an enormous innovative surge. It is therefore necessary to take into account future innovations in organism identification and susceptibility testing. New methods can influence procedures in the work process and drastically change the demands on capacity.

FUTURE PERSPECTIVES The degree of automation as well as the variability of the procedures that can be automated will continue to increase. Continued development will make the systems more flexible and result in products that can better accommodate the conditions of each individual lab. Automated technology is evolving quickly. It will soon include additional testing, e.g., direct testing of blood culture systems and molecular diagnostics. The primary objective of automating microbiology laboratories is to improve the quality and consistency of processes that suffer from high variability and are labor-intensive. The hope is that these technologies will allow the laboratory staff to concentrate on the processing of more technically demanding specimens. Software solutions for integrating POCT devices will further improve the analytic process. Lastly, digital image processing as an integral part of bacteriological clinical diagnostics will promote further development in telemedicine. In many centralized laboratory models, sample processing and plate reading may not take place in the same location. In such situations, quality standards can still be maintained in small labs or in remote areas by offering access to experienced personnel. Telemedicine can help counter the lack of skilled personnel in these areas. Further scientific evaluation of TLA could also facilitate an appraisal of its clinical relevance and its impact on patient care. It could reduce errors and improve the quality of diagnostic microbiology.

SUMMARY New advances in technology as well as staffing shortages are causing microbiologists to rethink laboratory design. It may be that the traditional laboratory model as we know it today will cease to exist in the near future. TLA will certainly become commonplace in high-volume laboratories and may eventually be found in smaller laboratories. Automation is unlikely to replace medical technologists, but it will change requirements in two important ways. First of all, manual processing of the plates will be replaced by digital imaging. Second, the efficiency of automation may cause more laboratories to adopt the 24-h culture-reading strategy. Clinical microbiology is changing at a rapid pace,

5. Laboratory Design and Workflow n 53

primarily due to a surge in technological advances. Laboratory designs will need to be more flexible to accommodate future developments. 16.

REFERENCES 1. US Department of Health and Human Services. 2009. Biosafety in Microbiological and Biomedical Laboratories, 5th ed. Centers for Disease Control and Prevention, Atlanta, GA. 2. Baselski V, Carey R, Clarridge J, Weissfeld A. 2005. Survey of Clinical Microbiology Laboratory Workloads, Productivity Rates and Staffing Vacancies. American Society for Microbiology, Washington, DC. 3. Jones JL, Lopez A, Wahlquist SP, Nadle J, Wilson M. 2004. Survey of clinical laboratory practices for parasitic diseases. Clin Infect Dis 38(Suppl 3):S198–S202. 4. Versalovic J, Funke G, Jorgensen J, Landry ML, Warnock D. 2011. Manual of Clinical Microbiology, 10th ed, vol 1. ASM Press, Washington, DC. 5. Campos J. 2012. Lean lab in action. MLO Med Lab Obs 44:25–29. 6. American Society of Clinical Pathologists. 1996. Board of Registry Study Guide. Clinical Laboratory Certification Examinations, 4th ed. American Society of Clinical Pathologists, Chicago, IL. 7. Wimmer JL, Long SW, Cernoch P, Land GA, Davis JR, Musser JM, Olsen RJ. 2012. Strategy for rapid identification and antibiotic susceptibility testing of gram-negative bacteria directly recovered from positive blood cultures using the Bruker MALDI Biotyper and the BD Phoenix system. J Clin Microbiol 50:2452–2454. 8. Tan KE, Ellis BC, Lee R, Stamper PD, Zhang SX, Carroll KC. 2012. Prospective evaluation of a matrix-assisted laser desorption ionization–time of flight mass spectrometry system in a hospital clinical microbiology laboratory for identification of bacteria and yeasts: a bench-by-bench study for assessing the impact on time to identification and costeffectiveness. J Clin Microbiol 50:3301–3308. 9. Patel R. 2013. Matrix-assisted laser desorption ionization– time of flight mass spectrometry in clinical microbiology. Clin Infect Dis 57:564–572. 10. Still M. 2013. MTRA, bitte melden. Dtsch Arzteblatt 110:105–106. 11. Mazzocato P, Savage C, Brommels M, Aronsson H, Thor J. 2010. Lean thinking in healthcare: a realist review of the literature. Qual Saf Health Care 19:376–382. 12. Riebling N, Condon S, Gopen D. 2004. Toward error free lab work. Six Sigma Forum Mag 4:6. 13. Rutledge J, Xu M, Simpson J. 2010. Application of the Toyota production system improves core laboratory operations. Am J Clin Pathol 133:24–31. 14. Carlson RO, Amirahmadi F, Hernandez JS. 2012. A primer on the cost of quality for improvement of laboratory and pathology specimen processes. Am J Clin Pathol 138:347–354. 15. Llopis MA, Trujillo G, Llovet MI, Tarres E, Ibarz M, Biosca C, Ruiz R, Kirchner MJ, Alvarez V, Busquets G, Domenech MV, Figueres C, Minchinela J, Pastor RM, Perich C, Ricos C, Sansalvador M, Palmada MS. 2011. Quality indicators and specifications for key analytical-

17.

18.

19. 20.

21. 22. 23.

24.

25.

26.

27.

28. 29.

30. 31.

32.

33.

34.

extranalytical processes in the clinical laboratory. Five years’ experience using the Six Sigma concept. Clin Chem Lab Med 49:463–470. Vanker N, van Wyk J, Zemlin AE, Erasmus RT. 2010. A Six Sigma approach to the rate and clinical effect of registration errors in a laboratory. J Clin Pathol 63:434– 437. Gras JM, Philippe M. 2007. Application of the Six Sigma concept in clinical laboratories: a review. Clin Chem Lab Med 45:789–796. Stankovic AK, Romeo P. 2007. The role of in vitro diagnostic companies in reducing laboratory error. Clin Chem Lab Med 45:781–788. Coskun A. 2007. Six Sigma and laboratory consultation. Clin Chem Lab Med 45:121–123. Persoon TJ, Zaleski S, Frerichs J. 2006. Improving preanalytic processes using the principles of lean production (Toyota production system). Am J Clin Pathol 125:16–25. Hawker CD. 2007. Laboratory automation: total and subtotal. Clin Lab Med 27:749–770, vi. Tilton RC, Ryan RW. 1978. Evaluation of an automated agar plate streaker. J Clin Microbiol 7:298–304. Mischnik A, Mieth M, Busch CJ, Hofer S, Zimmermann S. 2012. First evaluation of automated specimen inoculation for wound swab samples by use of the Previ Isola system compared to manual inoculation in a routine laboratory: finding a cost-effective and accurate approach. J Clin Microbiol 50:2732–2736. Bourbeau PP, Swartz BL. 2009. First evaluation of the WASP, a new automated microbiology plating instrument. J Clin Microbiol 47:1101–1106. Jones G, Matthews R, Cunningham R, Jenks P. 2011. Comparison of automated processing of flocked swabs with manual processing of fiber swabs for detection of nasal carriage of Staphylococcus aureus. J Clin Microbiol 49:2717– 2718. Glasson JH, Guthrie LH, Nielsen DJ, Bethell FA. 2008. Evaluation of an automated instrument for inoculating and spreading samples onto agar plates. J Clin Microbiol 46:1281–1284. Mulatero F, Bonnardel V, Micolaud C. 2011. The way forward for fast microbiology. Clin Microbiol Infect 17:661– 667. Bourbeau PP, Ledeboer NA. 2013. Automation in clinical microbiology. J Clin Microbiol 51:1658–1665. Dumitrescu O, Dauwalder O, Lina G. 2011. Present and future automation in bacteriology. Clin Microbiol Infect 17:649–650. Matthews S, Deutekom J. 2011. The future of diagnostic bacteriology. Clin Microbiol Infect 17:651–654. Van Horn KG, Audette CD, Sebeck D, Tucker KA. 2008. Comparison of the Copan ESwab system with two Amies agar swab transport systems for maintenance of microorganism viability. J Clin Microbiol 46:1655–1658. Fontana C, Favaro M, Limongi D, Pivonkova J, Favalli C. 2009. Comparison of the eSwab collection and transportation system to an Amies gel transystem for Gram stain of clinical specimens. BMC Res Notes 2:244. Greub G, Prod’hom G. 2011. Automation in clinical bacteriology: what system to choose? Clin Microbiol Infect 17:655–660. Garcia LS. 2010. Clinical Microbiology Procedures Handbook, 3rd ed. ASM Press, Washington, DC.

Molecular Microbiology* FREDERICK S. NOLTE

6 Since the publication of the 10th edition of this Manual, significant changes have occurred in the practice of diagnostic molecular microbiology. Nucleic acid amplification techniques are now commonly used to diagnose and manage patients with infectious diseases. The growth in the number of Food and Drug Administration (FDA)-cleared/approved test kits and analyte-specific reagents (ASRs) has facilitated the use of this technology in the clinical laboratory. Technological advances in nucleic acid amplification techniques, automation, nucleic acid sequencing, and multiplex analysis have reinvigorated the field and created new opportunities for growth. Simple, sample-in, answer-out molecular test systems are now available that can be deployed in a variety of laboratory and clinical settings. Molecular microbiology remains the leading area in molecular pathology in terms of both the numbers of tests performed and clinical relevance. Nucleic acid-based tests have reduced the dependency of the clinical microbiology laboratory on more traditional antigen detection and culture-based methods and created new opportunities for the laboratory to affect patient care. This chapter covers nucleic acid probes, signal and target amplification techniques, postamplification detection and analysis, clinical applications of these techniques, and the special challenges and opportunities that these techniques provide for the clinical laboratory. Applications of molecular methods used in epidemiological investigations, metagenomics, and new pathogen discovery are covered in chapters 10, 15, and 16, respectively.

The commonly used formats for probe hybridization include liquid-phase, solid-phase, and in situ hybridization. The leading method used in clinical microbiology laboratories is a liquid-phase hybridization protection assay (Hologic Gen-Probe, Inc., San Diego, CA). In this method, a singlestranded DNA (ssDNA) probe labeled with an acridinium ester is incubated with the target nucleic acid. Alkaline hydrolysis follows the hybridization step, and probe binding is measured in a luminometer after the addition of peroxides. For a positive sample, the acridinium ester on the bound probe is protected from hydrolysis and, upon the addition of peroxides, emits light. The hybridization protection assay can be completed in several hours and does not require removal of unbound single-stranded probe or isolation of probe-bound double-stranded sequences (1). In solid-phase hybridization, target nucleic acids are bound to nylon or nitrocellulose and are hybridized with a probe in solution (2). The unbound probe is washed away, and the bound probe is detected by means of fluorescence, luminescence, radioactivity, or color development. Although solid-phase hybridization is a powerful research tool, the length of time required and the complexity of the procedure limit its application in clinical practice. In situ hybridization is another type of solid-phase hybridization in which the nucleic acid is contained in tissues or cells that are affixed to microscope slides and is governed by the same basic principles described previously (3). In most clinical applications, formalin-fixed, paraffin-embedded tissue sections are used. The sensitivity of in situ hybridization is often limited by the accessibility of the target nucleic acid in the cells. In general, due to the poor analytical sensitivities of nonamplified-probe techniques, the application of these techniques to direct detection of pathogens in clinical specimens is limited to those situations in which the number of organisms is large. Such situations include cases of group A streptococcal pharyngitis and agents associated with vaginosis and vaginitis. These techniques are used most effectively in culture confirmation assays for mycobacteria and systemic dimorphic fungi. These culture confirmation tests have a positive effect on patient management by providing rapid and accurate detection of these slowly growing, often difficult-to-identify pathogens. Nucleic acid probes for direct detection of group A streptococci, Chlamydia trachomatis, and Neisseria gonorrhoeae are available from Hologic Gen-Probe. Probes for identification of Blastomyces dermatitidis, Coccidioides immitis, Histoplasma

NONAMPLIFIED NUCLEIC ACID PROBES Nucleic acid probes are segments of DNA or RNA labeled with radioisotopes, enzymes, or chemiluminescent reporter molecules that can bind to complementary nucleic acid sequences with high degrees of specificity. Although probes can range from 15 to thousands of nucleotides in size, synthetic oligonucleotides of 1,000 IU/ml at week 4 or 12 and/or detectable at week 24, then therapy should be discontinued. Currently, there is no clinical indication for viral resistance testing, but that may change as different classes of direct-acting antiviral agents are used for treatment. Quantitative tests for HIV-1 RNA are the standard of practice for guiding clinicians in initiating, monitoring, and changing antiretroviral therapy. Several commercially available HIV-1 viral load assays have been FDA approved, and guidelines for their use in clinical practice have been published (184). Viral load assays have also been used in monitoring response to therapy in patients chronically infected with HBV (185) and in predicting the risk for developing BK virus-associated nephropathy in renal transplant recipients (186). In organ transplant recipients, the persistence of CMV viral load after several weeks of antiviral therapy is associated with the development of resistance (187).

LABORATORY PRACTICE The unparalleled analytical sensitivity of nucleic acid amplification techniques coupled with their susceptibility to cross contamination presents unique challenges to the routine application of these techniques in the clinical laboratory. There are special concerns in the areas of specimen processing, workflow, quality assurance, and interpretation of test results. Additional information can be found in the CLSI documents MM3-A2, Molecular Diagnostic Methods for Infectious Diseases; Approved Guideline—2nd Edition (188); MM6A2, Quantitative Molecular Methods for Infectious Diseases; Approved Guideline—2nd Edition (142); MM13-A, Collection, Transport, Preparation, and Storage of Specimens and Samples for Molecular Methods; Approved Guideline (189); and MM19-A, Establishing Molecular Testing in Clinical Laboratory Environments; Approved Guideline (190).

Specimen Collection, Transport, and Processing Proper collection, transport, and processing of clinical specimens are essential to ensure reliable results from molecular

assays. Nucleic acid integrity must be maintained throughout these processes. Important issues to consider in specimen collection are the timing of specimen collection in relationship to disease state and the proper specimen type. Other factors that come into play include the use of the proper anticoagulant, transport and storage temperatures, and time to processing of the specimen. HIV-1 viral load testing is an example in which the proper conditions for specimen collection, transport, and processing have been well described and has provided insight into the importance of these factors. For HIV-1 viral load testing, the plasma needs to be separated from the cells within 6 h of collection to minimize degradation of RNA. Once the plasma has been separated, it can be stored at 4°C for several days, but −70°C is recommended for long-term storage (191). Most types of specimens are best stored at −20 to −70°C prior to processing. Molecular methods have several advantages over conventional culture with regard to specimen collection. It may be easier to maintain the integrity of nucleic acid than the viability of an organism. Molecular tests for the detection of C. trachomatis and N. gonorrhoeae are an example in which DNA is stable on dry cervical swabs for a week at room temperature or refrigeration temperatures, which is in stark contrast to the conditions required to maintain organism viability for culture. Nucleic acid persists in specimens after initiation of treatment (192, 193), thus allowing detection of a pathogen even though the organism can no longer be cultured. Also, due to the increased sensitivity of molecular assays, it may be possible to test a smaller volume of specimen or use a specimen that is collected using a less invasive method. The major goals of specimen processing are to release nucleic acid from the organism, maintain the integrity of the nucleic acid, render the sample noninfectious, remove inhibiting substances, and in some instances concentrate the specimen. These processes need to be balanced with minimizing manipulation of the specimen. Complex specimen processing methods are time-consuming and may lead to the loss of target nucleic acid or result in contamination between specimens. Care must be taken to avoid carrying over inhibitory substances, such as phenol or alcohol, from the nucleic acid isolation step to the amplification reaction. There are several general methods for nucleic acid extraction. Different methods may be used depending on whether the desire is to purify RNA or DNA or both. Another factor to consider when deciding on a nucleic acid extraction method is the type of pathogen sought. Some pathogens, such as viruses, can be very easy to lyse, while mycobacteria, Gram-positive bacteria, and fungi can be very difficult to lyse. Enzyme digestion, harsh lysis conditions, or mechanical disruption may be required to disrupt the cell walls of these organisms. DNA isolation methods often use detergents to solubilize the cell wall or membranes, a proteolytic enzyme (such as proteinase K) to digest proteins, and EDTA to chelate divalent cations needed for nuclease activity (194, 195). The lysate can be used directly in amplification assays, or additional steps may follow to purify the nucleic acid. These additional steps remove proteins and traces of organic solvents and concentrate the specimen. In order to successfully use a crude lysate, the target DNA must be present in a relatively high concentration and there must be minimal inhibitors of amplification in the sample. If these criteria are not met, additional purification steps should be used. Another commonly used method of nucleic acid isolation involves disruption of cells or organisms with the chaotropic

78

n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

agent guanidinium thiocyanate and a detergent (196). After a short incubation, the nucleic acid can be precipitated with isopropanol. Guanidinium thiocyanate denatures proteins and is also a strong inhibitor of ribonucleases, making it a very useful tool for RNA isolation, although it is also used for purification of DNA. The Boom extraction method is also based on the lysing and nuclease-inactivating properties of guanidinium thiocyanate but utilizes the acid-binding properties of silica or glass particles to purify nucleic acid (197). Detection of target organisms that are present in small numbers in a large-volume clinical sample requires that target organisms be concentrated to a detectable level. One way to accomplish this is to isolate the particular nucleic acid of interest by binding it to a solid phase, which allows the support, with the target bound to it, to be separated from the original sample. These techniques are referred to as target capture. Target capture techniques immobilize nucleic acids on magnetic beads by the use of a capture probe that attaches to the bead and to the target nucleic acid. A magnetic separation device is used to concentrate the target by drawing the magnetic beads to the sides of the sample tube, while the remainder of the sample is washed away and removed. Target capture techniques also remove materials in the sample that might otherwise interfere with amplification. Over the past several years, various manufacturers have developed commercially available reagents using one of these basic methods or a modification of these methods. Many of these methods rely on the use of spin column technology, are easy to use, and provide a rapid, reproducible method for purification of nucleic acid from a wide variety of clinical specimens. In recent years, further advances have been made with the introduction of magnetic silica particles that are coupled with instruments providing various degrees of automation, thus further simplifying nucleic acid extraction and purification. These reagents tend to be expensive, but the additional cost can be offset by labor savings. Laboratories are increasingly using automated systems for nucleic acid extraction, as they require less hands-on time, may reduce the risk of cross contamination between specimens, and provide more consistent yields. There are now many automated systems available for use in clinical laboratories; they should be thoroughly evaluated because not all isolate nucleic acids with the same efficiency and purity. The quality of the nucleic acid can have a significant impact on the performance of a molecular test. Tissue samples need to be disrupted prior to the nucleic acid extraction process. This can be accomplished by cutting the tissue into small pieces or mechanically homogenizing the tissue prior to proceeding with one of the above-described extraction methods. Preserved tissue specimens require removal of the paraffin with solvents and slicing into fine sections prior to processing. Removing inhibitors of amplification is a key function of the nucleic acid extraction process. Simple methods of nucleic acid extraction that involve boiling of the specimen have been used for relatively acellular specimens such as cerebrospinal fluid (CSF). Though the boiling method is fast and easy, there are problems with inhibitors of amplification in CSF that are not inactivated by boiling (198). The inhibition rate can be reduced to 10 kb, all of these systems generate between several hundred thousand and several billion short-read sequences between 100 and 800 bp long. Despite this similarity, there are nonetheless important differences in read length, sequence output, turnaround time, accuracy, error model, and cost (114–116). Each of these factors must be considered when selecting an appropriate sequencing strategy for specific objectives and applications. To further complicate matters, as NGS technologies continue to mature, the hardware, software, chemistries, and consumables are all subject to intense development and innovation, with improvements to sample throughput and data quality, and continual decreases in per-isolate sequencing costs. WGS represents the ultimate tool in molecular epidemiology, as it allows the identification of single genomic changes between two isolates. The technology has not yet been widely used for real-time surveillance and outbreak investigations at the community level, though it has proven its power in a few recent investigations in health care settings (117–119). WGS has been widely used to develop new diagnostic tests for known and emerging pathogens (120), and it has become an important tool in pathogen discovery and characterization and in studying pathogen evolution and transmission. A recent example is the analysis of the complete genome of the swine origin H1N1 strain that emerged in 2009, which showed that the virus had been circulating in humans for an extended period as the result of a single introduction. The new strain had genome segments derived from swine, human, and avian strains (121). Similarly, in the 2011 German outbreak of E. coli, WGS played a key role in understanding the determinants and modeling the evolutionary events that led to a hypervirulent strain (122, 123).

As the cost and complexity of sequencing continues to decline, the management and analysis of the massive volumes of output data have emerged as a significant challenge to the routine use of NGS for infectious disease surveillance, outbreak detection, and response. In order to be useful for public health intervention, raw whole-genome sequence data from suspected pathogens must be analyzed, assembled, interpreted, and compared in a timely fashion, so that action may be taken to prevent or minimize disease-associated morbidity and mortality. In the case of a small outbreak investigation, these data may represent tens of gigabytes of raw sequence, but for large molecular surveillance efforts, such as the PulseNet network for foodborne disease surveillance (124), the expected raw sequence output could easily surpass a hundred terabytes of raw genomic sequence data each year. The challenge of genomic “big data” will require the development of a dedicated informatics infrastructure to support NGS and other high-throughput laboratory technologies and the incorporation of genomics, bioinformatics, and data science skill sets by the public health workforce. Since most end users of the technology will be microbiologists in clinical and public health laboratories with limited bioinformatics skills, it is critical that user-friendly and intuitive bioinformatics tools be developed for most sequencingrelated surveillance activities. Equally important for global health is the development of clear and concise international consensus standards on sequence quality; data compression, storage, and transmission; sequence-associated metadata; and analytical methods for outbreak detection and response. Work to address these issues has already been initiated through the Global Microbial Identifier (GMI) network (http://www.globalmicrobialidentifier.org/), which consists of approximately 200 experts from at least 30 countries, including clinical, food, and public health microbiologists and virologists, bioinformaticians, epidemiologists, representatives from funding agencies, data hosting systems, and policymakers from academia, public health, industry, and governments. While most viral sequences can be directly compared from WGS data using multiple-sequence alignment, the analysis of most bacterial, fungal, and parasitic genomes is complicated by their larger relative sizes. While some WGSbased subtyping strategies for these organisms consider the entire genomic sequence, most focus on sets of specific markers or polymorphic regions within the genomic sequence. Several of the most common strategies are described below. A glossary of terms commonly used in bioinformatics is provided in Table 3.

Whole-Genome SNP Typing Whole-genome single nucleotide polymorphism (SNP) typing involves mapping sequence reads from each query (or test) isolate against a reference genome to identify and compare positions of single nucleotide variations. The reference sequence is typically selected from among closely related, finished genomes from public repositories or from a high-quality de novo assembly of an epidemiologically important internal reference isolate (e.g., an index case or putative source) in cases where an external reference sequence is either not available or not appropriate for the investigation. An example of the latter case is illustrated in Fig. 4. The initial pool of candidate SNPs can vary greatly in number between organisms according to their genome sizes, their complexity and plasticity, their population diversity, and the quality and coverage of mapped sequence reads. Candidate SNPs identified from each mapping are typically

142 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS TABLE 3

Definitions commonly used in bioinformatics

Term

Definition

Annotation

The identification and functional determination of genes and genetic elements within a sequence.

Genome assembly

Process by which many short DNA sequence fragments, such as those generated by next-generation sequencers, are reassembled into a representation of the original genomic sequence.

Contig

Contiguous consensus sequences derived from the assembly of many short, overlapping DNA fragments.

Coverage (read depth)

Avg number of reads representing a given nucleotide in the reconstructed sequence.

De novo assembly

Genomic assembly without an a priori reference sequence to inform the genomic structure.

High-confidence SNP

Single nucleotide polymorphism that has been verified using specific criteria, such as sequence coverage, sequence quality, and population and allelic frequency. While these parameters will vary depending on the context and application of the SNP calling, the typical minimum criteria are 10 to 20× coverage, >Q20 (see below for definition), and 100% frequency.

Indel

Insertion or deletion of nucleotides, which results in a net change of the total number of nucleotides.

k-mer

Oligonucleotide of length “k.”

N50 statistic

Commonly used as a rough assessment of genomic assemblies. It represents the contig length for which all contigs of that length or greater include at least half of the total base pair length of the sequence set.

Open reading frame

Uninterrupted reading frame in a given sequence that may be used for gene prediction.

Per-base sequence quality

The sequence quality score for each individual base position in a sequence. Typically, Phred scores are used, where Q = –10log(error probability). A Q30, for example, means a 1-in-1,000 likelihood of an incorrect base call at that position.

Read

A unit of continuous DNA sequence derived from target DNA.

Reference-mapped assembly

Assembly of a genome by aligning new sequence reads against a preexisting and similar reference sequence.

Suffix array

An array of all substring suffixes of a longer string. For example, the sequence ATGCATGC, represented as an array (A[i]), includes suffixes of ATGCATGC, ATGCATG, ATGCAT, ATGCA, ATGC, ATC, AT, and A.

filtered according to sequence quality; the position, genomic distribution, and functional impact of variant alleles; and population-level considerations, such as allelic frequency, genetic convergence, homoplasy, and linkage disequilibrium. With sufficient bioinformatics expertise and computational resources, the distillation from reference mappings to a minimum set of high-confidence, parsimoniously informative loci can be completed relatively quickly. Once validated, sets of high-confidence canonical SNPs can be used as targets for the analysis and subtyping of subsequent isolates or as the basis for high-throughput surveillance as-

says using real-time PCR or other lower-cost molecular technologies (125, 126). There are several circumstances in which SNP-based strain typing may perform suboptimally, primarily due to its dependence on consistent reference mapping. For some organisms, there are no circularized reference genomes necessitating the use of de novo assemblies as references. Even when a closed reference genome exists, genomic plasticity in the population may be sufficiently high in species such as Neisseria gonorrhoeae or Burkholderia that mapping to historical reference strains may be inefficient or result in

10. Molecular Epidemiology n

143

FIGURE 4 Heat map showing the SNPs among 10 STEC O157 isolates belonging to the same outbreak. In order to elucidate the genetic relationships among the outbreak isolates, high-quality SNPs were called against the outbreak reference strain (index case) that appears at the far left. A total of nine SNPs were detected among the outbreak isolates. doi:10.1128/9781555817381.ch10.f4

important gaps in genomic coverage. Similarly, in organisms where critical virulence and diversity factors are carried by plasmids, transposons, or other mobile genetic elements, information about important discriminatory loci may be absent from the available reference sequences. One strategy is to construct composite reference sequences from the microbial chromosome, with major plasmids or transposons of interest appended as a pseudoassembly. While this approach allows for the consideration of these extrachromosomal sequences, the limited stability of many mobile genetic elements and important differences in selective pressure between chromosomal and extrachromosomal alleles may present important problems. Some organisms also contain mutational hot spots (clustered mutations) from which SNP calls need to be assessed carefully, with the natural error rate of Taq polymerase kept in mind. A final consideration relates primarily to fungi and parasites, where the ploidy (number of sets of chromosomes) of the organism and the presence of complex or heterogenous alleles may impact the number and quality of high-confidence, discriminatory SNPs that are useful for subtyping and analysis.

WGS, MLST, and Binary Typing As the sequencing, assembly, and annotation of large numbers of microbial genomes become increasingly cost-effective and feasible, it is possible to define and query largescale MLST or binary typing schemes, which include dozens or even hundreds of different genes or sequences of interest. This approach can be applied to both microbial subtyping and characterization, as important differences in gene complement may be associated with the mobilization of plasmids, transposons, bacteriophages, or horizontal gene transfer from other organisms and may signal phenotypic differences in levels of virulence or antimicrobial susceptibility. Publicly available database schemas and software, such as the Bacterial Isolate Genome Sequence Database (BIGSdb) (http://pubmlst.org/software/database/bigsdb/), allow bacterial genomes to be catalogued and systematically

compared across many different MLST targets (127). Other measures, such as BLAST (Basic Local Alignment Search Tool) score ratios (BSR), provide a more general assessment of the functional similarity of two genomes, based on the pairwise comparison of all open reading frames (ORFs) (128). Although gene-based comparisons are emerging as an important strategy for strain typing and characterizing bacteria and other pathogens using WGS data, current limitations with short-read next-generation sequencing and bioinformatic analysis may impact their usefulness, particularly for large-scale comparisons. The reliance of these techniques on high-quality genomic assemblies and the consistent prediction or annotation of open reading frames present challenges in terms of the technical complexity and throughput of the analysis. Draft bacterial genomes may include hundreds of contigs, and the interpretation of complex MLST schemes will invariably be affected by breaks, duplications, or misassemblies of important sequences or reading frames. Differences in assembly or gene prediction algorithms may also impact the throughput of the assay and interpretation of results, and the development of standardized bioinformatic methods and workflows is critical, particularly as sequence databases scale to hundreds or thousands of genomes with many different alleles. Binary typing data for the presence or absence of genes or genetic variants may also be used to provide useful information for microbial characterization, particularly when used in conjunction with other strain typing methods. Interrogating sets of known resistance and virulence markers can provide important information during an outbreak investigation or response, and these features can often be used to support strain type identification or to correlate with transmission or patient outcomes.

k-mer Analysis Recently, a number of different k-mer-based approaches have emerged to analyze and compare large genomic data

144 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

sets in a computationally efficient manner. In general, these algorithms deconstruct genomic sequences or unassembled raw reads into large sets of k-mers, or oligonucleotides of length “k,” for comparison by means of suffix arrays. The k-mers are then sorted, deduplicated, and compared to other k-mer sequence sets to identify variations (SNPs and indels) that can be used for clustering and comparison (129). Unlike traditional SNP-mapping approaches, which take into consideration the genomic context of the variant position and the underlying sequence quality of the reads that support its identification, k-mer-based SNP analysis focuses on the putative SNP position and the 10 to 12 bases of surrounding sequence, without consideration of quality scores. These are important limitations, which may restrict the ability of k-mer-based methods to reliably detect closely spaced SNPs or indels and result in limited resilience against low sequence quality or sequencing errors. Even so, k-mer methods have significant advantages in terms of speed and implementation, since they do not require assembly or alignment of sequences and can be equally applied to wholegenome sequence sets and unassembled sets of short reads. k-mer-based algorithms continue to improve, and recent implementations for strain typing can assess horizontal gene transfer, annotate SNP positions based on a known reference, and accommodate missing alleles and evolutionary convergence, and they are easily adapted to new applications, such as the metagenomic analysis of complex samples (129). During outbreak investigations in particular, k-merbased SNP analysis is an excellent tool for initial genomic comparison and clustering, since it provides actionable strain typing information quickly and can be used to guide the application of more-resource-intensive downstream bioinformatics analyses, which may take several days to complete.

DNA Microarrays A DNA microarray is a molecular platform in which a few hundred to thousands of specific DNA oligonucleotides or short sequences (capture molecules) are bound to a matrix. The array is used to detect specific DNA sequences (target sequences) in a test sample of fluorescently labeled DNA, with or without amplification, by hybridization followed by detection of the array-bound test DNA. Two types of arrays, planar and liquid, are used. Analysis with planar microarrays is typically performed by deposition of DNA probes complementary to the genome targets of interest on a glass slide (solid matrix). Probes are typically synthetically produced oligonucleotides (9 to 100 bp) or PCR products (100 to 1,000 bp). They can target ORFs amplified from the sequenced reference isolate or short intergenic oligonucleotides based on the available sequence (130). Alternatively, PCR probes may be attached to microspheres that are internally dyed with two spectrally distinct fluorophores and combined to create a suspension array (liquid matrix), such as those seen with the Luminex platform (131). Overall, platforms developed may differ in a number of aspects, such as probe content, density, deposition technology, labeling, and hybridization of probes, as well as sample detection and analysis. An ORF-based DNA array can detect losses of entire ORFs but does not detect minor deletions, point mutations, deletions restricted to intergenic spaces, genetic rearrangements, deletions of homologous repetitive elements, or gene insertions. Short oligonucleotide arrays are more precise at detecting shorter nucleotide polymorphisms but require a larger number of probes at high cost. Such oligonucleotide arrays have been developed by companies such as Affymetrix (Santa Clara, CA), Operon

(Huntsville, AL), Agilent Technologies (Santa Clara, CA), and NimbleGen (Madison, WI). Insertions of genes compared with the sequenced reference strain cannot be detected in comparative genome hybridization DNA array analysis. This problem can be alleviated by adding nonredundant amplified sequences from several closely related organisms to the array (132). DNA microarrays represent an attractive platform for use in microbial identification and subtyping, since they allow for rapid and accurate analysis of large numbers of different DNA molecules. Besides its remarkable resolution, DNA array analysis unveils the genetic region responsible for diversity, often allowing phenotypic predictions at the same time. In addition, due to the basic design of microarrays (i.e., detection of signal associated with sample DNA bound to fixed DNA probes), the positional variation associated with gel-based methods, such as PFGE, is eliminated and the analysis of results is easier to automate and standardize. Limitations of DNA array technology include the initial high cost for the synthesis and spotting of target-specific probes and for the fluorophores used to label the reactions. Ambiguities in the interpretation of the ratios of hybridization and cross-hybridization to analogous targets are also important limitations of the technique (133). The interlaboratory reproducibility of DNA array-based methods remains unclear, since multilaboratory validation studies are largely missing (134). Even though planar arrays allow for the analysis of hundreds of thousands of targets at the same time, sample throughput is an obvious bottleneck of the technology. If the total number of probes can be limited to a few hundred, suspension arrays offer a higher sample throughput platform with lower cost. Currently, the Luminex xMAP system (Luminex Corp., Austin, TX) allows for up to 100 different probes to be analyzed simultaneously in a single well of a 96-well plate. The main application for DNA arrays is pathogen identification. One approach is to amplify more-universal genes (e.g., 16S rRNA, 18S rRNA, 23S rRNA) and to screen for pathogen-specific polymorphisms (135, 136). However, universal PCR can be challenging, since commercial PCR reagents are frequently contaminated with trace amounts of bacterial DNA. Additionally, since the detection is based on pathogen-specific point mutations in these genes, naturally occurring variants may not all be represented on the array, resulting in false-negative detection events. An alternative strategy for coupling PCR and DNA array detection is to use multiplex PCR to amplify a number of discrete, pathogen-specific genetic markers. The ability to construct high-density microarrays with multiple probe sequences makes it relatively trivial to include multiple markers for specific pathogens and thus permit greater validity of a positive detection event (137). The disadvantage of this approach is that there is a practical limit to the number of primer sets that can be included in the PCR. Instead of coupling PCR amplification and DNA array detection, applying direct detection to extracted DNA or RNA is achievable when the amount of target is abundant (138). The primary goal in this case is to avoid amplification biases associated with PCR. Several microarrays have been developed for rapid identification of viruses, including the ViroChip and the GreeneChip (139–141). These arrays have the capacity to detect a large number of viral pathogens and can quickly identify the etiologic agent in unknown source outbreaks (142, 143). In some cases, the arrays are designed to detect specific groups of viruses, such as respiratory pathogens (144–146). The most comprehensive pathogen detection

10. Molecular Epidemiology n

arrays reported to date were designed by a team at the Lawrence Livermore National Laboratory. The Lawrence Livermore microbial detection array (LLMDA) can test over 6,000 viruses and 15,000 bacteria as well as fungi and protozoa (147, 148). The array contains both “discovery” probes, which match conserved genome regions that are unique to a taxonomic family or a subfamily but shared by species within that family, facilitating detection of novel species within a family, and “census” probes, which target highly variable regions unique to an individual species or a strain and can hence be used for forensic purposes. Microarrays have also been used to genotype viral and bacterial agents based on SNPs occurring on specific sequence regions. In many cases, though, whole-genome sequence analysis may be the more cost-effective means of obtaining a viral or bacterial genotype. Nevertheless, there are numerous examples of the use of microarrays for viral genotyping, including human rotavirus (149), human and avian influenza viruses (150, 151), hepatitis B virus (152), HIV (153), varicella-zoster virus (154), measles virus (155), human papillomavirus (156, 157), polioviruses (158), and noroviruses (142, 143). Similarly, microarrays have been utilized for bacterial subtyping by molecular serotyping (159), SNP analysis (160, 161), and binary typing (162) and for detection of the presence or absence of antimicrobial resistance genes (163, 164) or specific mutations conferring antimicrobial resistance (165, 166). Microarrays, besides being used for pathogen identification and genotyping, have been used for resequencing the complete genomes of viral agents (167, 168) and partial bacterial genomes (169, 170); however, the development of next-generation sequencing methods has made arraybased sequencing impractical.

Mass Spectrometry In recent years, matrix-associated laser desorption ionization–time of flight (MALDI-TOF) MS systems have become increasingly common in clinical and public health microbiology laboratories throughout the world. In contrast to earlier generations of mass spectrometry equipment, which were complex, costly, and difficult to maintain, these newer benchtop MALDI-TOF systems offer simple and standardized workflows for many genus- and species-level microbial identification tasks, with high throughput, accurate and fast results, and a low overall per-test cost (171). While a few proof-of-concept studies have demonstrated the feasibility of MALDI-TOF for strain- and subspecies-level identification of select organisms (172–175), standardized or generalizable methods have yet to be developed, and in most cases, the current level of resolution of MALTI-TOF, when used alone, remains at the genus or species level. Attempts have been made by some commercial companies to combine mass spectrometry with other technologies and typing approaches. The PLEX-ID (Abbott Laboratories, Abbott Park, IL) system uses PCR and reverse transcription-PCR (RTPCR) electrospray ionization mass spectrometry to identify and/or type viruses, bacteria, and fungi. For most bacterial species, the level of resolution is generally equivalent to that of MLST, with the inclusion of other important multiplexed targets for further confirmation or characterization (176). Another system, the Agilent MassCode PCR system (Agilent Technologies), uses highly multiplexed PCR panels and identifies/subtypes microorganisms through the detection and measurement of cleavable mass tags using atmospheric-pressure, chemical-ionization, quadrupole mass spectrometry (177). Other platforms, such as the Sequenom MassArray (Sequenom Inc., San Diego, CA), use PCR

145

MALDI-TOF to interrogate thousands of SNPs through a single primer base extension and can be used for highresolution SNP-based strain typing using standardized sets of canonical targets (178). However, these applications remain under active development and in most cases to do not currently provide the discriminatory power needed for epidemiologically relevant high-resolution strain typing.

APPLICATIONS Molecular Surveillance Molecular Serotyping Serotyping is a subtyping method in which specific antigens on the surfaces of microorganisms are detected by antigenantibody reactions using diagnostic antisera. The method is definitive but only moderately discriminatory. It has nevertheless proven to be extremely useful for epidemiologic classification of a number of pathogens and has for this reason been a mainstay in the surveillance of different infectious diseases for more than 70 years. It is a cumbersome method that requires the availability of high-quality antisera and substantial expertise to perform the assay and interpret the results. More convenient, rapid, and reliable molecular alternatives to serotyping are being developed. These molecular serotyping methods are designed to generate data that are in close or complete agreement with conventional serotyping in order not to lose the connection to the historical data. In the previous sections, molecular flagellin typing of Campylobacter (flaA typing), M serotyping (emm typing) of GAS, and capsular (cps) typing of pneumococci have been mentioned. In the genus Salmonella, more than 2,500 serotypes are currently recognized (179). Two types of molecular serotyping approaches have been used; in one, a molecular subtyping method is correlated with the serotype, and in the second, the genes acrually encoding the serotype are targeted directly. In this latter case, for O antigens the genes are within the rfb gene cluster, and for the flagellar antigens the fliC and fljB genes encode phases 1 and 2, respectively. Some examples of subtyping methods that correlate with serotyping are MLST, PFGE, ribotyping, and rep-PCR (180); the correlation between subtyping methods is never perfect, and ideally, serotypes determined this way should be confirmed by traditional serotyping or a molecular serotyping method targeting the serotype-encoding genes. Regarding the latter, the U.S. Centers for Disease Control and Prevention (CDC) has developed an assay that reliably identifies the vast majority of the serotypes involved in human infections using a combination of PCR amplification of the rfb genes, fliC, and fljB, with detection of the amplicons using a liquid microarray (159, 181). This assay was recently commercialized by Luminex. With the advances in sequencing technology, it is also becoming increasingly cost-efficient to determine the serotype directly from the sequences of the serotype-encoding genes (182). The methods targeting rfb, fliC, and fljB are directly compatible with the Kauffman-White scheme, which contains the phenotypic descriptions of all Salmonella serotypes (179), and the results will not need to be confirmed by traditional serotyping.

Cluster Detection/Outbreak Investigations Subtyping methods that are used for community-wide routine surveillance to detect disease case clusters and to support outbreak investigations need to be rapid, highly dis-

146 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

FIGURE 5 PFGE profiles of two STEC O157 strains, namely, the most common sporadic pattern, EXHX01.0047, and the outbreak-associated pattern, EXHX01.1264, and their distribution during the 2002 outbreak period. doi:10.1128/9781555817381.ch10.f5

criminatory, reproducible, and amenable to library subtyping so that trends in the occurrence of the organisms that they target may be followed easily. Often subtyping results are the only information available to determine whether an outbreak investigation should be initiated. In this situation, the case definition needs to be very specific; i.e., the discriminatory power of the subtyping method needs to be high in order not to include patients that are not associated with the outbreak as cases, which would confuse the epidemiological investigation. The molecular methods that are most suitable for this purpose are PFGE, MLVA, AFLP (when used in one laboratory), and highly discriminatory DNA array and gene-sequencing methods. These methods may be used in conjunction with less discriminatory traditional and molecular methods, e.g., biotyping, serotyping, phage typing, antibiograms, and/or plasmid profiling. PulseNet, the national molecular subtyping network for foodborne disease surveillance, is a prime example of a network performing real-time surveillance of infectious diseases at the community level (124). In a subtyping network, the choice of method is less important than that all participants use the same method, as long as the preferred method has sufficient performance characteristics. In PulseNet, the preferred subtyping method is PFGE, and only isolates with indistinguishable patterns are considered when clusters are being detected. An outbreak of STEC O157:H7 that occurred in the summer of 2002 was caused by a strain with a pattern that differed from the most common pattern in the PulseNet database by just one band (Fig. 5A). Fortyfour case patients with the outbreak pattern were identified, but during the outbreak period, 41 isolates with the highly similar most common pattern were also detected (Fig. 5B); in case-control or other interview studies, the inclusion of these 41 patients infected with isolates displaying the common pattern as cases would have completely obscured the association with the vehicle, ground beef from one particular plant (183). As more supporting epidemiological information is gathered, it may be possible to amend the microbiological case definition. In the above-mentioned outbreak, two cases with epidemiological links to the outbreak were added to the list of case patients, even though

they were infected with a strain displaying the most common variant pattern. The requirements for methods used for surveillance and investigation of outbreaks in hospitals and other institutions may be different from the ones used in the community. The setting is much smaller in scale and usually attended by a single laboratory, and the outbreaks are often caused by pathogens transmitted from person to person and/or with pathogens that persist in the environment. Strains that are involved in outbreaks transmitted between patients or persistently from the environment often evolve during the outbreak, and variant subtypes may be present at the same time. This was the basis for the much-cited “Tenover criteria” for interpretation of PFGE results (184); they state that strains may be related if they differ from each other by up to six restriction bands, corresponding to two genetic events. Obviously, these criteria do not apply to point source outbreaks. The methods that are used for community-wide surveillance may also be used for these nosocomial outbreaks, and methods that do not show good interlaboratory reproducibility may be used as long as the intralaboratory reproducibility is sufficient. AFLP has excellent intralaboratory reproducibility, and even rep-PCR may be used for short-term surveillance and to delineate outbreaks that have been detected by other means, e.g., by phenotypic methods (185–187). The combination of molecular epidemiologic techniques and standard case classification and reporting provides a very sensitive means of describing the transmission pathways of many viruses. In particular, analysis of the sequence data can help to confirm the source of a virus or suggest a source for cases in which the source was unknown. Molecular data can be used to establish epidemiologic links, or lack thereof, between various cases and outbreaks. Molecular techniques have been particularly useful for the study of outbreaks of foodborne viruses, such as norovirus (188). In these cases, sequence data were used to identify the source and to trace the transport and distribution of contaminated food (189, 190). For highly infectious viruses, such as measles virus, transmission can occur anywhere. Molecular techniques have been used to identify the source of the infections when standard case investigations have failed to identify a source (191).

Source Attribution Source attribution is the epidemiological science that studies the relative contributions of different sources to the burden of infectious diseases. Strictly speaking, outbreak investigations are source attribution analyses, but the term is more commonly used in a broader sense encompassing both outbreak-related and sporadic disease. Molecular methods may be used to study the geographic distribution or sources and spread of microbial strains, and this is discussed in a section below. Source attribution using microbiological methods is gaining increasing attention, especially in the study of foodborne disease, to estimate where in the food production chain the largest public health problems originate. The decision makers use these estimates to prioritize where the control efforts can be spent most efficiently. A subtyping method that is used for source attribution creates subtypes that are distributed unevenly between the different possible sources of human infection. When this knowledge is combined with information about the distribution of the same subtypes in sick humans and, if available, the human rates of exposure to each source, it is possible to estimate the relative contri-

10. Molecular Epidemiology n

bution of each source to the burden of human disease (192). This approach was introduced in the United Kingdom to attribute the sources of Campylobacter jejuni and Campylobacter coli to predominantly chicken meat, with ruminants being another important source (193), and has since been used in many countries to attribute the sources of campylobacteriosis and to guide and to demonstrate the effect of intervention measures (194, 195). The limitation of the model is that it will attribute only to sources for which there are data available; i.e., if no isolates are included from a potential major source, no estimates for this source will be generated. Subtyping methods used for source attribution do not need to be as discriminatory or reproducible as those used for outbreak investigations, as long as they clearly differentiate isolates from different sources. Choosing a subtyping method with an unnecessarily high discriminatory power will increase the complexity of the analysis of the data set. In regard to Campylobacter, PFGE is the gold standard for outbreak investigations, whereas the less discriminatory method MLST works well for attribution. Theoretically, subtyping methods that provide phylogenetic information are more useful for source attribution than methods that do not provide this information, since the data collected from each source are likely not to be all-inclusive. Even if some subtyping information is missing, the data may still be useful, since strains that are related phylogenetically to each other are more likely to originate from the same source.

Dynamics of Infectious Disease Pathogen Evolution Molecular techniques have made enormous contributions to our understanding of the evolution of pathogens, but a detailed description of these studies is beyond the scope of this chapter. Molecular techniques have been used to monitor the distribution of circulating strains of various pathogens, to monitor the stability of antigenic sites that are important for diagnostics and as vaccine targets, and to assess susceptibility to antimicrobial drugs. Improvements in computational molecular biology resources and new analytical methods such as BEAST (Bayesian Evolutionary Analysis and Sampling Trees) (196) incorporate sample date and allow the evolutionary dynamics of a population to be inferred from sequence data. These approaches are called phylodynamics (197). With these approaches, sequence information from infectious agents can be used to infer the transmission patterns of that agent over time. Analysis of hepatitis C virus sequences from Egypt suggested an exponential increase in the number of cases in 1930 and 1955, when unsterile needles were used to distribute an antischistosomal therapy (198). Similar studies documented the episodic transmission of HIV into London during the late 1990s (199). More recently, these methods were used to investigate the origins of the swine origin influenza A H1N1 (2009) viruses that were associated with the influenza pandemic in 2009 and 2010 (200).

Geographic Spread The ability of a molecular technique to identify the source of an infectious agent can be applied on a global scale, and the information can be used to monitor the spread of the pathogen. The sensitivity of this approach depends on the level of surveillance activity and the availability of a global database of genetic information. Sequence analysis was used to identify the source of the West Nile virus that was intro-

147

duced into the United States in 1999 and to track the spread of this lineage of virus across the entire country (201, 202), as well as to track the global spread of various genetic variants of HIV (203).

Sustained Transmission in Areas That Are Destined for Elimination Molecular surveillance for viral diseases that are prevented by vaccination is especially beneficial when it is possible to observe the change in viral genotypes over time in a particular country or region. This information, when analyzed in conjunction with standard epidemiologic data, has helped to document the interruption of transmission of endemic viruses and provides a means to measure the success of vaccination programs. Molecular characterization of measles viruses has provided a valuable tool for measuring the effectiveness of measles control programs. In general, three patterns of measles genotype distribution have been described. In countries that still have endemic transmission of measles, the majority of cases are caused by several endemic genotypes that are distributed geographically. In these cases, multiple cocirculating lineages within the endemic genotype or genotypes are present. In countries that have eliminated measles, the small numbers of cases are caused by a number of different genotypes that reflect various sources of imported virus and suggest the lack of sustained transmission of an endemic genotype or genotypes. The third pattern occurs in countries or regions that have had very good measles control but are experiencing an increase in the numbers of susceptible individuals because of failure to maintain high rates of vaccination coverage. In this situation, reintroduction of measles usually results in a large outbreak associated with a single genotype of virus with nearly identical sequences (191, 204– 206).

Pathogen Discovery/Identification Molecular methods are sometimes crucial for the discovery of new pathogens, especially viral pathogens, and to determine if a pathogen is truly new or has arisen from known pathogens through recombination or other genetic events. For example, coronaviruses have the ability to recombine with other coronaviruses as well as to incorporate cellular genes into their genomes (207). The initial analysis of the complete genome sequence of the coronavirus associated with severe acute respiratory syndrome (SARS-CoV) confirmed that the virus was a novel coronavirus and not a recombinant between previously described coronaviruses (208, 209). In 2012, rapid WGS of a novel coronavirus causing acute respiratory syndrome in humans (HCoV EMC or MERS-CoV) showed that the new virus was a new variant of Betacoronavirus that was closely related to a bat coronavirus and more distantly related to SARS-CoV (210–212).

Vaccination Issues Live attenuated vaccines are used to control a number of viral diseases. In some cases, these vaccines can cause symptoms that are similar to those caused by infection with the wild-type virus. When the risk of exposure to disease is very low, these vaccine reactions can be identified by temporal association with vaccination. However, when vaccination is used as part of an outbreak response, it may be difficult to distinguish vaccine reactions from symptoms caused by the wild-type virus. Since serologic techniques usually lack the sensitivity to distinguish between antibodies directed at the vaccine or the wild-type strain, genetic characteriza-

148 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

tion of the viruses is usually the only means available to clearly classify the case as vaccine associated or due to a wild-type infection. Of course, this approach requires that appropriate specimens for virologic detection be collected from the suspected cases and that sequence information be available for the vaccine strains. The availability of molecular methods that can clearly identify vaccine reactions, as described above, has contributed to our knowledge about the safety of live attenuated vaccines. In addition, molecular techniques now play an important role in the postlicensure evaluation of live attenuated viral vaccines. Here, genetic characterization is used to monitor the stability of vaccines in the field and to clearly identify vaccine reactions. In one example, PCR and RFLP analysis were used to monitor the safety of varicella-zoster vaccine by classifying vesicular rash or zoster cases as being associated with wild-type varicella-zoster virus of the Oka vaccine strain (213). Additionally, some viruses have the capacity to recombine or reassort their genomes, and in these cases, WGS is an important component of molecular epidemiologic studies. The live attenuated strains used in the WHO polio eradication program replicate in the human gut and can be excreted for several weeks after immunization. If the attenuating mutations in the vaccine strains revert, the vaccine may cause vaccine-associated paralytic poliomyelitis in vaccinees or result in transmissible, neurovirulent, circulating vaccine-derived poliovirus strains which have been associated with outbreaks of poliomyelitis (214, 215). The vaccine-derived polioviruses often contain mosaic genomes that result from recombination between the vaccine strains and other lineages of poliovirus or other enteroviruses (215, 216), and these variants can be detected only by full-genome sequencing. In some cases, data from molecular surveillance of viral and bacterial pathogens are used to decide on the most appropriate formulation for vaccines. The geographic distribution of strains may affect the efficacy of the vaccines if the genetic changes are accompanied by changes in antigenicity. Therefore, for agents with variable antigenic properties (e.g., rotavirus and influenza virus), careful monitoring of the strains associated with cases in vaccinated populations is necessary (217–219).

Forensic Microbiology Sometimes infectious disease incidents are criminally investigated if there is a suspicion that they may be the result of deliberate actions or criminal gross negligence. HIVinfected individuals have been imprisoned for spreading the virus intentionally (220) or through negligence (221), and anthrax has been spread through the mail (86, 222). Numerous outbreaks of foodborne infections occur each year, and some have been the result of intentional contamination of the food supply (223). Forensic molecular epidemiology comes into play when regulators and laymen, judges, and lawyers have to decide if an infection (or infections) is a result of criminal action. In principle, criminal microbiological investigations are not different from outbreak investigations, except that in criminal investigations, the link between the infection(s) and its (their) source(s) needs to be proven beyond reasonable doubt. In outbreak investigations, the precautionary principle is relatively often used when deciding on public health actions, since the aim of an outbreak investigation is to stop the outbreak from spreading as soon as possible. This is in contrast to criminal investigations, where the precautionary principle is not used since an offender is presumed innocent until proven otherwise. Subtyping methods that may quantify the probability that

two microbial strains are related are therefore preferred over methods that do not yield direct quantifiable information. Non-target-specific methods are therefore not ideal for criminal investigations because DNA fragments of the same size may have completely different sequence contents; e.g., it is not possible from the PFGE pattern to tell if two isolates are identical, hence the use of the term “indistinguishable” for PFGE patterns that cannot be differentiated from each other. In contrast, differences observed with a target-specific method may be quantifiable if the prevalence of the detected alleles is known in the relevant microbial populations.

SUBTYPING METHOD SELECTION, VALIDATION, AND DATA INTERPRETATION Method Selection All too often, it appears that the selection of subtyping methods is guided only by convenience criteria, in particular, the accessibility of a method to the laboratory. This has led to numerous studies in which an inappropriate method was used to study an organism (1). When selecting a subtyping method, one should first of all consider the epidemiological context in which it is going to be used; i.e., what is the question that the typing data should answer? A method that is appropriate for long-term surveillance is most likely not appropriate for a short-term investigation of a nosocomial outbreak. Careful consideration should also be given for the genetic makeup of the target organisms, including the clonality, mode of transmission, and outbreak potential.

Factors That Impact the “Clonality” of a Given Population The clonal relatedness of isolates is manifested by their display of a significantly higher level of similarity in their genotype and/or phenotype than can be expected from randomly sampled and epidemiologically unrelated isolates of the same species (1). The simplest explanation for a genetically monomorphic pathogen is that the population size of the ancestors of all extant organisms was so strongly reduced during a recent bottleneck that genetic diversity was abolished. One possibility that can result in such a bottleneck is a crucial genetic event that happened only once, such as a change in ecological niche due to the acquisition of two plasmids by the progenitor of Yersinia pestis (224). In the case of STEC O157, it has been suggested that clonality can be explained by source-sink evolution dynamics (225– 227). According to this theory, mutations in microbes that reside mainly in reservoirs (the source, e.g., cattle) in which they are not pathogens confer on a small subset a phenotype that results in injury to an accidental host (the sink, e.g., humans). Alternatively, it has been argued that a strong selective advantage conferred by a mutation enabled certain strains of STEC O157 to flourish in cattle, making them more available for spillover into humans (227). On the other hand, some host-restricted pathogens, such as S. enterica serotype Typhi, are under relatively little selection pressure from their host or environment and therefore do not diversify through point mutations, recombination, or acquisition of new sequences (228). Finally, a sampling bias may also contribute to the appearance of population clonality. Sampling from one part of the phylogenetic tree will overlook much of the variation present in the population and collapse all isolates outside the studied population into a single type (229). In order to be able to adequately discriminate clonal organisms, subtyping methods targeting fast-evolving genetic elements, such as VNTRs or SNPs, are usually preferred (230, 231). With some organisms, optimal results

10. Molecular Epidemiology n

may be obtained by using multiple techniques, e.g., PFGE and VNTR analysis (232).

Method Validation Once a proper method or approach has been selected, it must be subjected to the highest scrutiny possible to ensure that it meets the criteria discussed at the beginning of this chapter. This includes validation using a population of strains and isolates that originate from the epidemiological context to which the method is going to be applied. Techniques that will be applied in local investigations need to be validated on a strain collection from the same locality, whereas methods that will be used for community-wide surveillance need to be validated on a collection of strains reflecting the diversity in the whole community. When a subtyping method that has been validated for use in one given epidemiological context is to be used in a different one, it may be necessary to validate it in the new context before it is implemented. This supplementing validation may not need to be as thorough as the original validation, depending on how similar the two contexts are. Unlike methods developed in isolation (for use by one laboratory), methods to be used in multiple laboratories for the generation of data archived in reference libraries must be particularly carefully tested, evaluated, and validated. All methods need to go through four phases of validation: initial development, internal validation, external validation, and, finally, postimplementation evaluation. The first two phases often overlap. Additionally, before the new method is implemented, a quality assurance and quality control (QA/QC) program needs to be established. The goal of the initial development is to identify the optimal conditions or parameters to ensure that the protocol is robust and reproducible and generates highly discriminatory and epidemiologically concordant data on all strains. Ten to 50 isolates that represent the diversity in the study population at large is typically used in this phase. During the internal validation, the method is tested by individuals not involved in the method development in order to ascertain the robustness of the protocol in the hands of laboratorians with no prior experience with it. The panel of test isolates is expanded to include the full genetic diversity of the study population and should contain both sporadic and outbreak-related isolates in order to test the true discriminatory power and the epidemiological concordance of the method. Duplicate isolates of the same strain and multiple isolates from single-source outbreaks need to be included to evaluate the reproducibility and stability of the method. The test panel will usually contain 250 to 500 isolates. The isolates need to be selected from a collection of strains with a known subtype if a gold standard subtyping method exists for the organism in order to be able to evaluate the performance of the new method against this gold standard. During the external validation, the robustness and portability of the method is further tested typically by 5 to 10 external partner laboratories, ideally with different levels of subtyping expertise and access to different types or brands of equipment and reagents. The assay is evaluated using 10 to 50 isolates selected by the laboratory that developed the method. The interlaboratory reproducibility of the method is also assessed during this phase. Sometimes the external validation is further expanded to include a prospective or retrospective testing of up to 50 isolates from each laboratory’s own collection. Following the successful completion of these phases, the method may be implemented. However, even after its imple-

149

mentation, the performance of the assay needs to be assessed on a regular basis to detect problems not identified during the initial validation and to assess emerging situations, such as the impact of introduction of new brands of reagents that may have become commercially available.

Quality Assurance/Quality Control A quality assurance program needs to be in place before any molecular subtyping technique is implemented to ensure consistently high quality and reproducibility of the data generated. At a minimum, subtyping should be performed only by personnel trained in working with the procedure; a written standard operating procedure should be in place, and a strain with a well-established stable subtype should be included in all experiments in order to detect procedural failures. For library subtyping systems involving multiple laboratories, participation in an external quality assessment (EQA) program that includes an initial certification and annual proficiency testing is mandatory.

Data Interpretation When interpreting subtyping results, one must consider the epidemiological context and all other available data, such as associated demographic and other epidemiological information, and other subtyping information, e.g., biochemical reaction profiles, serotype, phage type, antimicrobial susceptibility profiles, and the presence of virulence factors; reliance on a single parameter for characterization should be avoided whenever possible. Knowledge of the subtyping method, including the quality of the data, the diversity of the organism, and the history of the subtypes encountered, should also be considered (233).

Quality of the Data Even when a carefully standardized procedure is followed, artifacts may occur, which may lead to erroneous conclusions about relationships between profiles. It is therefore important to know the nature of these artifacts in order to recognize and correct them. In this, the role of the database curator is extremely important. In PFGE, the artifacts include, among other things, ghost bands caused by incomplete restriction and subtle differences in band resolution (one thick band versus two thinner bands) (233). In PCRbased methods, such as RAPD and rep-PCR, differences in band intensities are a huge problem, and in MLVA, PCR, and fragment analysis artifacts, such as minus-A and stutter peaks and fluorescence carryover, can confuse data interpretation (234). Also, Sanger sequence trace files should be routinely checked for quality either manually or by using software (e.g., Phred/Phrap, http://www.phrap.org). Most major sequence databases require the submission of the raw sequence trace files from the laboratory when a new allele type is proposed (235). The quality of the next-generation WGS data can be assessed using various quality metrics that are based on the raw reads (e.g., per-base sequence quality, sequence length distribution, sequence duplication level, and sequence coverage) or assemblies (e.g., N50 and the number of contigs) and can be generated by using either the software built into the sequencing system or external software packages. It is important to recognize that the quality of the genome assemblies reflects the quality of the sequencing technology used but also of the analysis software employed for assembly and annotation (236).

Diversity of the Organism Optimal interpretation of the differences (or lack thereof) between subtypes of two isolates depends largely on the

150 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

variability of the organism being typed. Large databases should provide sufficient data to make reasonable determinations of diversity. If an organism displays little diversity, one should be cautious in assuming that closely related patterns, or even indistinguishable patterns, indicate a high likelihood that they originate from a common source. In this case, additional data from other typing methods and other available information should be considered. If the organism shows substantial diversity, one must still consider whether there are clonal subpopulations within a nonclonal organism. On the other hand, when an organism demonstrates extreme variability, any pattern matches may be significant.

Epidemiological Context If the epidemiological setting from which the isolates are derived appears to be a point source outbreak without continued transmission, only very minor differences are likely to be observed, because the outbreak strain has very little time to undergo genetic changes. In contrast, when there is ongoing transmission, such as prolonged hospital or community outbreaks, with strains being passed from person to person, more variability should be expected. Additionally, the amount of variation seen during an outbreak will depend on the stability of the genetic markers targeted by the typing method. Fast-evolving genetic markers, such as VNTRs, tend to change slightly even during shorter outbreaks (90). During investigations of known outbreaks, it is fairly easy and often helpful to designate patterns that differ slightly from the primary outbreak pattern as subpatterns or variants. However, when performing surveillance for cluster/outbreak detection, accepting such variations may mislead the epidemiological investigation (183), especially if one of the variants represents a common pattern. It is also important to consider how the laboratory results fit together with the epidemiologic and environmental investigations. The fact that common subtypes exist for many organisms demonstrates that indistinguishable subtypes alone do not unequivocally prove an epidemiologic connection. It is always possible that two individuals were infected with the same strain from different sources. On the other hand, clearly differing subtypes do not prove that isolates are epidemiologically unrelated, since multiple strains can be associated with the same outbreak (237) and some strains contain genetic elements, e.g., prophages, which are incorporated into the genome in an unstable manner and therefore may affect the subtype of different colonies picked from the same pure culture of the organism (238).

LIBRARIES FOR MOLECULAR EPIDEMIOLOGY International travel, migration, and the food trade are the main factors that have contributed to the worldwide spread of microbes. Therefore, the need for international databases with standardized type nomenclature and information on epidemiologically relevant strains has emerged. Building such databases relies upon standardization of typing methods and on regular ring trials for all participating laboratories to guarantee consistently comparable data.

Strain Catalogues Strain catalogue databases are mainly public access websites with limited access components in some of them. Their main function is to standardize subtype nomenclature and facilitate easy data sharing. Most of them are curated but typically contain minimal epidemiological or demographic

information about the strains and are therefore not very useful for real-time epidemiological surveillance. The laboratories that contribute data to strain catalogues use protocols that vary in their levels of standardization from none (MLVA) to medium (spa typing). The best-known examples of strain libraries are the three MLST databases that are hosted on Web servers located at Imperial College London (http://www.mlst.net) (235), Oxford University (http://pubmlst.org) (239), and the Institut Pasteur (http://www.pasteur.fr/mlst). On http:// www.mlst.net and the Institut Pasteur websites, speciesspecific information, including limited epidemiological data, can be accessed on each of the species websites, whereas http://pubmlst.org software enables multiple client databases to query a single species-specific profile database, so that the information on isolates can be kept private for security or confidentiality reasons. Each species-specific database holds the sequences of all known alleles at each of the MLST loci and through the curator assigns new allele numbers and sequence types. The websites also provide clustering tools to explore the relationship of the query strain with other strains in the database. SeqNet (http://www.seqnet.org/), coordinated by the Münster University Hospital and Robert Koch Institute in Wernigerode, Germany, is an example of a non-MLSTbased strain catalogue. SeqNet is an initiative of 60 national reference laboratories and university laboratories from 29 European countries to establish a network of sequence-based typing of microbial pathogens (240). SeqNet currently has only a database for S. aureus spa types through the curated spa server. Unlike the three MLST databases, SeqNet has a QA/QC aspect built in it in the form of a one-time certification and annual ring trials. The ccrB typing tool (http://www.ccrbtyping.net) is a public online resource for storage and automatic analysis of MRSA ccrB sequences (241). The user’s sequence is assigned to an allele based on 100% homology with an existing allele or to a new allele if a homology between 90% and 100% is found to any of the available alleles. In the case of a new allele, the most similar allele is indicated. Based on the allele assignment, a prediction of the ccrAB allotype and SCCmec type is also provided. International databases for MLVA data have also been recently established. MLVAbank (http://mlva.u-psud.fr), hosted by the University of Orsay, France, has both public and private databases available for a few organisms (94). The public databases have been derived from published data, sometimes by merging publications from different groups. Since the VNTR markers used by different groups are not always the same because of the current lack of standardization, different data sets may be available for different sets of isolates. Since different sets of primers are sometimes used for the same marker by different groups and since there are sizing discrepancies between different capillary electrophoresis platforms, the data are not always directly comparable even though they are uploaded to MLVAbank as repeat copy numbers. The TB database (http://www.tbdb.org) is an integrated database for tuberculosis (TB) research that houses annotated genome sequence data for several M. tuberculosis strains, microarray and RT-PCR expression data from in vitro experiments, and TB-infected tissues (243). Experimental data may be deposited into the database by any TB researcher prior to publication, providing prepublication access to tools for the analysis, annotation, visualization, and sharing of data. The data are then made public at the author’s request or following publication. The database

10. Molecular Epidemiology n

curators also actively search the literature for publications containing relevant TB or host microarray data and then obtain the raw data from the researchers and load them into the database.

Surveillance Databases Surveillance databases are restricted-access curated databases for real-time sharing of subtyping data and detailed demographic information associated with the strains. The main function of surveillance databases is to rapidly detect and define clusters of disease in order to initiate and support epidemiological investigations that are aimed at tracing the source and limiting the scope of outbreaks. Laboratories contributing data to surveillance databases typically are required to follow highly standardized protocols and comply with an extensive QA/QC program to ensure the reproducibility and the high quality of data. PulseNet, the national and international surveillance network for foodborne disease, is the most successful example of a surveillance database. PulseNet USA was established in 1996 (20), and similar networks later followed in Canada, the Asia Pacific region, Latin America, the Middle East, and Europe, and the latest is in Africa (244) (www.pulse netinternational.org). The participating laboratories perform PFGE on all foodborne organisms that they receive from clinical and food specimens in real time. The generated patterns are analyzed locally using highly customized software and uploaded to the central databases of each region via the Internet. The database managers confirm the quality of the patterns, name them according to a standardized scheme, and compare them against the patterns submitted to the database within the previous 60 days (120 days for Listeria). The epidemiologists are alerted if a new cluster of indistinguishable isolates is detected. Until now, much of the work in PulseNet International has focused on establishing the infrastructure of the network, but eventually each regional network will be able to log on to the server of any other international network, query the databases for matches to pathogen subtypes of interest, and, if matches are found, access the epidemiologic information on matching isolate patterns (244). Such collaboration is already taking place between PulseNet USA and PulseNet Canada. Other PulseNet-like surveillance databases include the CaliciNet network in the United States (245) and the Foodborne Viruses in Europe (FBVE) network (246), dealing mainly with norovirus. The virological networks employ primarily DNA sequence analysis. A number of virus-specific, curated databases have been established. These can be identified by an Internet search, though many may have restricted access.

CONCLUSIONS AND FUTURE TRENDS Molecular methods have improved our understanding of the epidemiology of infectious diseases and the well-known emerging or reemerging pathogens causing them during the past 4 decades. Molecular methods have been used not only to subtype and otherwise characterize the pathogens following their culture but also to identify and detect nonculturable or slowly growing organisms (247–249). The emergence and increased use of a plethora of cultureindependent diagnostic tests (250, 251) in clinical laboratories in the last few years has resulted in decreased availability of cultures for further characterization in public health laboratories. This trend may place culture-dependent surveillance networks such as PulseNet in serious jeopardy. However, with the rapid evolution of next-generation se-

151

quencing technologies, including metagenomics (252), it is to be expected that these methods will be used to an increasing extent for rapid nonculture diagnostics of infectious diseases directly from a clinical sample, providing detection, virulence characterization, and subtyping of the organisms at the same time, thereby creating the potential for detection of public health events, e.g., outbreaks, in real time, with no delays caused by isolating and growing the organisms as pure cultures. Before this can be accomplished, some formidable challenges need to be tackled, such as building comprehensive sequence reference libraries for metagenomics containing a representative diversity not only of pathogenic microbes but also of the human microbiome (253). Additionally, a significant investment in public health informatics infrastructure is needed to ensure that the users of the data are alerted to and know how to use the growing information flowing from clinical laboratories.

REFERENCES 1. van Belkum A, Tassios PT, Dijkshoorn L, Haeggman S, Cookson B, Fry NK, Fussing V, Green J, Feil E, GernerSmidt P, Brisse S, Struelens M. 2007. Guidelines for the validation and application of typing methods for use in bacterial epidemiology. Clin Microbiol Infect 13(Suppl 3):1– 46. 2. Hunter PR. 1990. Reproducibility and indices of discriminatory power of microbial typing methods. J Clin Microbiol 28:1903–1905. 3. Shannon CE, Weaver W. 1949. The Mathematical Theory of Communication. University of Illinois Press, UrbanaChampaign, IL. 4. Kado CI, Liu ST. 1981. Rapid procedure for detection and isolation of large and small plasmids. J Bacteriol 145:1365– 1373. 5. Meyers JA, Sanchez D, Elwell LP, Falkow S. 1976. Simple agarose gel electrophoretic method for the identification and characterization of plasmid deoxyribonucleic acid. J Bacteriol 127:1529–1537. 6. Mayer LW. 1988. Use of plasmid profiles in epidemiologic surveillance of disease outbreaks and in tracing the transmission of antibiotic resistance. Clin Microbiol Rev 1:228– 243. 7. Tofteland S, Naseer U, Lislevand JH, Sundsfjord A, Samuelsen O. 2013. A long-term low-frequency hospital outbreak of KPC-producing Klebsiella pneumoniae involving intergenus plasmid diffusion and a persisting environmental reservoir. PLoS One 8:e59015. 8. Pezzoli L, Elson R, Little CL, Yip H, Fisher I, Yishai R, Anis E, Valinsky L, Biggerstaff M, Patel N, Mather H, Brown DJ, Coia JE, van Pelt W, Nielsen EM, Ethelberg S, de Pinna E, Hampton MD, Peters T, Threlfall J. 2008. Packed with Salmonella—investigation of an international outbreak of Salmonella Senftenberg infection linked to contamination of prepacked basil in 2007. Foodborne Pathog Dis 5:661–668. 9. Haraoui L-P, Lévesque S, Lefebvre B, Blanchette R, Tomkinson M, Mataseje L, Mulvey MR, Miller MA. 2013. Polyclonal outbreak of KPC-3-producing Enterobacter cloacae at a single hospital in Montréal, Québec, Canada. J Clin Microbiol 51:2406–2408. 10. Kobayashi S, Wada A, Shibasaki S, Annaka M, Higuchi H, Adachi K, Mori N, Ishikawa T, Masuda Y, Watanabe H, Yamamoto N, Yamaoka S, Inamatsu T. 2009. Spread of a large plasmid carrying the cpe gene and the tcp locus amongst Clostridium perfringens isolates from nosocomial outbreaks and sporadic cases of gastroenteritis in a geriatric hospital. Epidemiol Infect 137:108–113. 11. Buchman TG, Roizman B, Adams G, Stover BH. 1978. Restriction endonuclease fingerprinting of herpes simplex

152 n

12.

13.

14.

15.

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27. 28.

29.

30.

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

virus DNA: a novel epidemiological tool applied to a nosocomial outbreak. J Infect Dis 138:488–498. Kaper JB, Bradford HB, Roberts NC, Falkow S. 1982. Molecular epidemiology of Vibrio cholerae in the U.S. Gulf Coast. J Clin Microbiol 16:129–134. Scherer S, Stevens DA. 1987. Application of DNA typing methods to epidemiology and taxonomy of Candida species. J Clin Microbiol 25:675–679. Tungpradabkul S, Panyim S, Wilairat P, Yuthavong Y. 1983. Analysis of DNA from various species and strains of malaria parasites by restriction endonuclease fingerprinting. Comp Biochem Physiol B Biochem Mol Biol 74:481– 485. Goering RV, Ribot EM, Gerner-Smidt P. 2009. Pulsedfield gel electrophoresis: laboratory and epidemiologic considerations for interpretation of data, p 167–177. In Tenover FC (ed), Molecular Microbiology: Diagnostic Principles and Practice, 2nd ed. ASM Press, Washington, DC. Schwartz DC, Cantor CR. 1984. Separation of yeast chromosome-sized DNAs by pulsed field gradient gel electrophoresis. Cell 37:67–75. Carle GF, Frank M, Olson MV. 1986. Electrophoretic separations of large DNA molecules by periodic inversion of the electric field. Science 232:65–68. Carle GF, Olson MV. 1984. Separation of chromosomal DNA molecules from yeast by orthogonal-field-alternation gel electrophoresis. Nucleic Acids Res 12:5647–5664. Chu G, Vollrath D, Dacis RW. 1986. Separation of large DNA molecules by contour-clamped homogeneous electric fields. Anal Biochem 234:1582–1585. Swaminathan B, Barrett TJ, Hunter SB, Tauxe RV, the CDC PulseNet Task Force. 2001. PulseNet: the molecular subtyping network for foodborne bacterial disease surveillance, United States. Emerg Infect Dis 7:382–389. van Embden JD, Cave MD, Crawford JT, Dale JW, Eisenach KD, Gicquel B, Hermans P, Martin C, McAdam R, Shinnick TM, Small PM. 1993. Strain identification of Mycobacterium tuberculosis by DNA fingerprinting: recommendations for a standardized methodology. J Clin Microbiol 31:406–409. Southern E. 1975. Detection of specific sequences among DNA fragments separated by gel electrophoresis. J Mol Biol 98:503–517. Cave MD, Eisenach KD, McDermott PF, Bates JH, Crawford JT. 1991. IS6110: conservation of sequence in the Mycobacterium tuberculosis complex and its utilization in DNA fingerprinting. Mol Cell Probes 5:73–80. Githui WA, Wilson SM, Drobniewski FA. 1999. Specificity of IS6110-based DNA fingerprinting and diagnostic techniques for Mycobacterium tuberculosis complex. J Clin Microbiol 37:1224–1226. Aston C, Mishra B, Schwartz DC. 1999. Optical mapping and its potential for large-scale sequencing projects. Trends Biotechnol 17:297–302. Schwartz DC, Li X, Hernandez LI, Ramnarain SP, Huff EJ, Wang YK. 1993. Ordered restriction maps of Saccharomyces cerevisiae chromosomes constructed by optical mapping. Science 262:110–114. Miller JM. 2013. Whole-genome mapping: a new paradigm in strain-typing technology. J Clin Microbiol 51:1066–1070. Onmus-Leone F, Hang J, Clifford RJ, Yang Y, Riley MC, Kuschner RA, Waterman PE, Lesho EP. 2013. Enhanced de novo assembly of high throughput pyrosequencing data using whole genome mapping. PLoS One 8:e61762. Riley MC, Kirkup BC Jr, Johnson JD, Lesho EP, Ockenhouse CF. 2011. Rapid whole genome optical mapping of Plasmodium falciparum. Malar J 10:252. Saunders MP, Wu G, Abuoun M, Pan Z, Anjum M, Woodward MJ. 2011. Optical genetic mapping defines regions of chromosomal variation in serovars of S. enterica

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

subsp. enterica of concern for human and animal health. Epidemiol Infect 139:1065–1074. Shukla SK, Pantrangi M, Stahl B, Briska AM, Stemper ME, Wagner TK, Zentz EB, Callister SM, Lovrich SD, Henkhaus JK, Dykes CW. 2012. Comparative wholegenome mapping to determine Staphylococcus aureus genome size, virulence motifs, and clonality. J Clin Microbiol 50:3526–3533. Welsh J, McClelland M. 1990. Fingerprinting genomes using PCR with arbitrary primers. Nucleic Acids Res 18:7213–7227. Williams JGK, Kubelik AR, Livak KJ, Rafalski JA, Tingey SV. 1990. DNA polymorphisms amplified by arbitrary primers are useful as genetic markers. Nucleic Acids Res 17:6531–6535. Meunier JR, Grimont PAD. 1993. Factors affecting reproducibility of random amplified polymorphic DNA fingerprinting. Res Microbiol 144:373–379. Stern MJ, Ames GF, Smith NH, Robinson EC, Higgins CF. 1984. Repetitive extragenic palindromic sequences: a major component of the bacterial genome. Cell 37:1015– 1026. Versalovic J, Koeuth T, Lupski JR. 1991. Distribution of repetitive DNA sequences in eubacteria and application to fingerprinting of bacterial genomes. Nucleic Acids Res 19:6823–6831. Sharples GJ, Lloyd RG. 1990. A novel repeated DNA sequence located in the intergenic regions of bacterial chromosomes. Nucleic Acids Res 18:6503–6508. Martin B, Humbert O, Camara M, Guenzi E, Walker J, Mitchell T, Andrew P, Prudhomme M, Alloing G, Hakenbeck R, Morrison DA, Boulnois GJ, Claverys JP. 1992. A highly conserved repeated DNA element located in the chromosome of Streptococcus pneumoniae. Nucleic Acids Res 20:3479–3483. Deplano A, Schuermans A, Van Eldere J, Witte W, Meugnier H, Etienne J, Grundmann H, Jonas D, Noordhoek GT, Dijkstra J, van Belkum A, van Leeuwen W, Tassios PT, Legakis NJ, van der Zee A, Bergmans A, Blanc DS, Tenover FC, Cookson BC, O’Neil G, Struelens MJ. 2000. Multicenter evaluation of epidemiological typing of methicillin-resistant Staphylococcus aureus strains by repetitiveelement PCR analysis. The European Study Group on Epidemiological Markers of the ESCMID. J Clin Microbiol 38:3527–3533. Rasschaert G, Houf K, Imberechts H, Grijspeerdt K, De Zutter L, Heyndrickx M. 2005. Comparison of five repetitive-sequence-based PCR typing methods for molecular discrimination of Salmonella enterica isolates. J Clin Microbiol 43:3615–3623. Healy M, Huong J, Bittner T, Lising M, Frye S, Raza S, Schrock R, Manry J, Renwick A, Nieto R, Woods C, Versalovic J, Lupski JR. 2005. Microbial DNA typing by automated repetitive-sequence-based PCR. J Clin Microbiol 43:199–207. Amoureux L, Bador J, Siebor E, Taillefumier N, Fanton A, Neuwirth C. 2013. Epidemiology and resistance of Achromobacter xylosoxidans from cystic fibrosis patients in Dijon, Burgundy: first French data. J Cyst Fibros 12:170– 176. Czaban SL, Sacha P, Wieczorek P, Kłosowska W, Krawczyk M, Ojdana D, Poniatowski B, Tryniszewska E. 2013. Rep-PCR genotyping of infectious Acinetobacter spp. strains from patients treated in intensive care unit of emergency department (ICU of ED)—preliminary report. Adv Med Sci 58:164. Verroken A, Bauraing C, Deplano A, Bogaerts P, Huang D, Wauters G, Glupczynski Y. 2013. Epidemiological investigation of a nosocomial outbreak of multidrug-resistant Corynebacterium striatum at one Belgian university hospital. Clin Microbiol Infect 20:44–50.

10. Molecular Epidemiology n 45. Bouchet V, Huot H, Goldstein R. 2008. Molecular genetic basis of ribotyping. Clin Microbiol Rev 21:262–273. 46. Bidet P, Barbut F, Lalande V, Burghoffer B, Petit JC. 1999. Development of a new PCR-ribotyping method for Clostridium difficile based on ribosomal RNA gene sequencing. FEMS Microbiol Lett 175:261–266. 47. Cartwright CP, Stock F, Beekmann SE, Williams EC, Gill VJ. 1995. PCR amplification of rRNA intergenic spacer regions as a method for epidemiologic typing of Clostridium difficile. J Clin Microbiol 33:184–187. 48. Vos P, Hogers R, Bleeker M, Reijans M, van de Lee T, Hornes M, Frijters A, Pot J, Peleman J, Kuiper M, Zabeau M. 1995. AFLP: a new technique for DNA fingerprinting. Nucleic Acids Res 23:4407–4414. 49. Koeleman JG, Stoof J, Biesmans DJ, Savelkoul PH, Vandenbroucke-Grauls CM. 1998. Comparison of amplified ribosomal DNA restriction analysis, random amplified polymorphic DNA analysis, and amplified fragment length polymorphism fingerprinting for identification of Acinetobacter genomic species and typing of Acinetobacter baumannii. J Clin Microbiol 36:2522–2529. 50. Fry NK, Afshar B, Visca P, Jonas D, Duncan J, Nebuloso E, Underwood A, Harrison TG. 2005. Assessment of fluorescent amplified fragment length polymorphism analysis for epidemiological genotyping of Legionella pneumophila serogroup 1. Clin Microbiol Infect 11:704–712. 51. Manning SD, Motiwala AS, Springman AC, Qi W, Lacher DW, Ouellette LM, Mladonicky JM, Somsel P, Rudrik JT, Dietrich SE, Zhang W, Swaminathan B, Alland D, Whittam TS. 2008. Variation in virulence among clades of Escherichia coli O157:H7 associated with disease outbreaks. Proc Natl Acad Sci U S A 105:4868–4873. 52. Persson S, Olsen KE, Ethelberg S, Scheutz F. 2007. Subtyping method for Escherichia coli Shiga toxin (verocytotoxin) 2 variants and correlations to clinical manifestations. J Clin Microbiol 45:2020–2024. 53. Sugimoto N, Shima K, Hinenoya A, Asakura M, Matsuhisa A, Watanabe H, Yamasaki S. 2011. Evaluation of a PCR-restriction fragment length polymorphism (PCR– RFLP) assay for molecular epidemiological study of Shiga toxin-producing Escherichia coli. J Vet Med Sci 73:859–867. 54. Tien N, You BJ, Chang HL, Lin HS, Lee CY, Chung TC, Lu JJ, Chang CC. 2012. Comparison of genospecies and antimicrobial resistance profiles of isolates in the Acinetobacter calcoaceticus-Acinetobacter baumannii complex from various clinical specimens. Antimicrob Agents Chemother 56:6267–6271. 55. De Baere T, de Mendonca R, Claeys G, Verschraegen G, Mijs W, Verhelst R, Rottiers S, Van Simaey L, De Ganck C, Vaneechoutte M. 2002. Evaluation of amplified rDNA restriction analysis (ARDRA) for the identification of cultured mycobacteria in a diagnostic laboratory. BMC Microbiol 2:4. 56. Dauga C, Zabrovskaia A, Grimont PAD. 1998. Restriction fragment length polymorphism analysis of some flagellin genes of Salmonella enterica. J Clin Microbiol 36:2835– 2843. 57. Fields PI, Blom K, Hughes HJ, Helsel LO, Feng P, Swaminathan B. 1997. Molecular characterization of the gene encoding H antigen in Escherichia coli and development of a PCR-restriction fragment length polymorphism test for identification of E. coli O157:H7 and O157:NM. J Clin Microbiol 35:1066–1070. 58. Nachamkin I, Bohachick K, Patton CM. 1993. Flagellin gene typing of Campylobacter jejuni by restriction fragment length polymorphism analysis. J Clin Microbiol 31:1531– 1536. 59. Acke E, McGill K, Lawlor A, Jones BR, Fanning S, Whyte P. 2010. Genetic diversity among Campylobacter jejuni isolates from pets in Ireland. Vet Rec 166:102–106.

153

60. Magnusson SH, Guethmundsdottir S, Reynisson E, Runarsson AR, Harethardottir H, Gunnarson E, Georgsson F, Reiersen J, Marteinsson VT. 2011. Comparison of Campylobacter jejuni isolates from human, food, veterinary and environmental sources in Iceland using PFGE, MLST and fla-SVR sequencing. J Appl Microbiol 111:971–981. 61. Maiden MC, Bygraves JA, Feil E, Morelli G, Russell JE, Urwin R, Zhang Q, Zhou J, Zurth K, Caugant DA, Feavers IM, Achtman M, Spratt BG. 1998. Multilocus sequence typing: a portable approach to the identification of clones within populations of pathogenic microorganisms. Proc Natl Acad Sci USA 95:3140–3145. 62. Massire C, Gertz RE Jr, Svoboda P, Levert K, Reed MS, Pohl J, Kreft R, Li F, White N, Ranken R, Blyn LB, Ecker DJ, Sampath R, Beall B. 2012. Concurrent serotyping and genotyping of pneumococci by use of PCR and electrospray ionization mass spectrometry. J Clin Microbiol 50:2018– 2025. 63. Feil EJ, Li BC, Aanensen DM, Hanage WP, Spratt BG. 2004. eBURST: inferring patterns of evolutionary descent among clusters of related bacterial genotypes from multilocus sequence typing data. J Bacteriol 186:1518–1530. 64. Noller AC, McEllistrem MC, Stine OC, Morris JG Jr, Boxrud DJ, Dixon B, Harrison LH. 2003. Multilocus sequence typing reveals a lack of diversity among Escherichia coli O157:H7 isolates that are distinct by pulsed-field gel electrophoresis. J Clin Microbiol 41:675–679. 65. Zhang W, Jayarao BM, Knabel SJ. 2004. Multi-virulencelocus sequence typing of Listeria monocytogenes. Appl Environ Microbiol 70:913–920. 66. Bilek N, Martin IM, Bell G, Kinghorn GR, Ison CA, Spratt BG. 2007. Concordance between Neisseria gonorrhoeae genotypes recovered from known sexual contacts. J Clin Microbiol 45:3564–3567. 67. Wick LM, Qi W, Lacher DW, Whittam TS. 2005. Evolution of genomic content in the stepwise emergence of Escherichia coli O157:H7. J Bacteriol 187:1783–1791. 68. Odds FC, Jacobsen MD. 2008. Multilocus sequence typing of pathogenic Candida species. Eukaryot Cell 7:1075–1084. 69. Beall B, Facklam R, Thompson T. 1996. Sequencing emmspecific PCR products for routine and accurate typing of group A streptococci. J Clin Microbiol 34:953–958. 70. Facklam RF, Martin DR, Lovgren M, Johnson DR, Efstratiou A, Thompson TA, Gowan S, Kriz P, Tyrrell GJ, Kaplan E, Beall B. 2002. Extension of the Lancefield classification for group A streptococci by addition of 22 new M protein gene sequence types from clinical isolates: emm103 to emm124. Clin Infect Dis 34:28–38. 71. Leung MH, Bryson K, Freystatter K, Pichon B, Edwards G, Charalambous BM, Gillespie SH. 2012. Sequetyping: serotyping Streptococcus pneumoniae by a single PCR sequencing strategy. J Clin Microbiol 50:2419–2427. 72. Pai R, Gertz RE, Beall B. 2006. Sequential multiplex PCR approach for determining capsular serotypes of Streptococcus pneumoniae isolates. J Clin Microbiol 44:124–131. 73. Turlej A, Hryniewicz W, Empel J. 2011. Staphylococcal cassette chromosome mec (Sccmec) classification and typing methods: an overview. Pol J Microbiol 60:95–103. 74. International Working Group on the Classification of Staphylococcal Cassette Chromosome Elements (WGSCC). 2009. Classification of staphylococcal cassette chromosome mec (SCCmec): guidelines for reporting novel SCCmec elements. Antimicrob Agents Chemother 53:4961– 4967. 75. David MZ, Daum RS. 2010. Community-associated methicillin-resistant Staphylococcus aureus: epidemiology and clinical consequences of an emerging epidemic. Clin Microbiol Rev 23:616–687. 76. Domingo E. 2007. Virus evolution, p 389–422. In Fields BN, Knipe DM, Howley PM (ed), Fields’ Virology, 5th

154 n

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

89.

90.

91.

92.

93.

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

ed, vol 1. Wolters Kluwer Health/Lippincott Williams & Wilkins, Philadelphia, PA. Nathanson N. 2007. Epidemiology, p 423–443. In Fields BN, Knipe DM, Howley PM (ed), Fields’ Virology, 5th ed, vol 1. Wolters Kluwer Health/Lippincott Williams & Wilkins, Philadelphia, PA. Kashi Y, King D, Soller M. 1997. Simple sequence repeats as a source of quantitative genetic variation. Trends Genet 13:74–78. van Belkum A, Scherer S, van Alphen L, Verbrugh H. 1998. Short-sequence DNA repeats in prokaryotic genomes. Microbiol Mol Biol Rev 62:275–293. Torres-Cruz J, van der Woude MW. 2003. Slipped-strand mispairing can function as a phase variation mechanism in Escherichia coli. J Bacteriol 185:6990–6994. van Belkum A, Scherer S, van Leeuwen W, Willemse D, van Alphen L, Verbrugh H. 1997. Variable number of tandem repeats in clinical strains of Haemophilus influenzae. Infect Immun 65:5017–5027. Shopsin B, Gomez M, Montgomery SO, Smith DH, Waddington M, Dodge DE, Bost DA, Riehman M, Naidich S, Kreiswirth BN. 1999. Evaluation of protein A gene polymorphic region DNA sequencing for typing of Staphylococcus aureus strains. J Clin Microbiol 37:3556–3563. Keim P, Price LB, Klevytska AM, Smith KL, Schupp JM, Okinaka R, Jackson PJ, Hugh-Jones ME. 2000. Multiplelocus variable-number tandem repeat analysis reveals genetic relationships within Bacillus anthracis. J Bacteriol 182:2928–2936. Grissa I, Bouchon P, Pourcel C, Vergnaud G. 2008. Online resources for bacterial micro-evolution studies using MLVA or CRISPR typing. Biochimie 90:660–668. Lindstedt BA, Vardund T, Aas L, Kapperud G. 2004. Multiple-locus variable-number tandem-repeats analysis of Salmonella enterica subsp. enterica serovar Typhimurium using PCR multiplexing and multicolor capillary electrophoresis. J Microbiol Methods 59:163–172. Hoffmaster AR, Fitzgerald CC, Ribot E, Mayer LW, Popovic T. 2002. Molecular subtyping of Bacillus anthracis and the 2001 bioterrorism-associated anthrax outbreak, United States. Emerg Infect Dis 8:1111–1116. Dyet KH, Turbitt E, Carter PE. 2011. Multiple-locus variable-number tandem-repeat analysis for discriminating within Salmonella enterica serovar Typhimurium definitive types and investigation of outbreaks. Epidemiol Infect 139:1050–1059. Torpdahl M, Sorensen G, Lindstedt BA, Nielsen EM. 2007. Tandem repeat analysis for surveillance of human Salmonella Typhimurium infections. Emerg Infect Dis 13:388–395. Hopkins KL, Maguire C, Best E, Liebana E, Threlfall EJ. 2007. Stability of multiple-locus variable-number tandem repeats in Salmonella enterica serovar Typhimurium. J Clin Microbiol 45:3058–3061. Hyytia-Trees E, Smole SC, Fields PA, Swaminathan B, Ribot EM. 2006. Second generation subtyping: a proposed PulseNet protocol for multiple-locus variable-number tandem repeat analysis of Shiga toxin-producing Escherichia coli O157 (STEC O157). Foodborne Pathog Dis 3:118–131. Hyytia-Trees E, Lafon P, Vauterin P, Ribot EM. 2010. Multilaboratory validation study of standardized multiplelocus variable-number tandem repeat analysis protocol for Shiga toxin-producing Escherichia coli O157: a novel approach to normalize fragment size data between capillary electrophoresis platforms. Foodborne Pathog Dis 7:129–136. Nadon CA, Trees E, Ng LK, Møller Nielsen E, Reimer A, Maxwell N, Kubota KA, Gerner-Smidt P. 2013. Development and application of MLVA methods as a tool for inter-laboratory surveillance. Euro Surveill 18(35):20565. De Santis R, Ancora M, De Massis F, Ciammaruconi A, Zilli K, Di Giannatale E, Pittiglio V, Fillo S, Lista F.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

106.

107.

2013. Molecular strain typing of Brucella abortus isolates from Italy by two VNTR allele sizing technologies. Mol. Biotechnol 55:101–110. Gorge O, Lopez S, Hilaire V, Lisanti O, Ramisse V, Vergnaud G. 2008. Selection and validation of a multilocus variable-number tandem-repeat analysis panel for typing Shigella spp. J Clin Microbiol 46:1026–1036. Lobersli I, Haugum K, Lindstedt BA. 2012. Rapid and high resolution genotyping of all Escherichia coli serotypes using 10 genomic repeat-containing loci. J Microbiol Methods 88:134–139. Ishino Y, Shinagawa H, Makino K, Amemura M, Nakata A. 1987. Nucleotide sequence of the iap gene, responsible for alkaline phosphatase isozyme conversion in Escherichia coli, and identification of the gene product. J Bacteriol 169:5429–5433. Grissa I, Vergnaud G, Pourcel C. 2008. CRISPRcompar: a website to compare clustered regularly interspaced short palindromic repeats. Nucleic Acids Res 36:W145–W148. Barrangou R, Fremaux C, Deveau H, Richards M, Boyaval P, Moineau S, Romero DA, Horvath P. 2007. CRISPR provides acquired resistance against viruses in prokaryotes. Science 315:1709–1712. Sorek R, Kunin V, Hugenholtz P. 2008. CRISPR—a widespread system that provides acquired resistance against phages in bacteria and archaea. Nat Rev Microbiol 6:181–186. Pourcel C, Salvignol G, Vergnaud G. 2005. CRISPR elements in Yersinia pestis acquire new repeats by preferential uptake of bacteriophage DNA, and provide additional tools for evolutionary studies. Microbiology 151:653–663. Grissa I, Vergnaud G, Pourcel C. 2007. CRISPRFinder: a web tool to identify clustered regularly interspaced short palindromic repeats. Nucleic Acids Res 35:W52–W57. Groenen PMA, Bunschoten AE, Soolingen D, van Embden JD. 1993. Nature of DNA polymorphism in the direct repeat cluster of Mycobacterium tuberculosis; application for strain differentiation by a novel typing method. Mol Microbiol 10:1057–1065. Fabre L, Zhang J, Guigon G, Le Hello S, Guibert V, Accou-Demartin M, de Romans S, Lim C, Roux C, Passet V, Diancourt L, Guibourdenche M, IssenhuthJeanjean S, Achtman M, Brisse S, Sola C, Weill FX. 2012. CRISPR typing and subtyping for improved laboratory surveillance of Salmonella infections. PLoS One 7:e36995. Song EJ, Jeong HJ, Lee SM, Kim CM, Song ES, Park YK, Bai GH, Lee EY, Chang CL. 2007. A DNA chipbased spoligotyping method for the strain identification of Mycobacterium tuberculosis isolates. J Microbiol Methods 68:430–433. Liu F, Barrangou R, Gerner-Smidt P, Ribot EM, Knabel SJ, Dudley EG. 2011. Novel virulence gene and clustered regularly interspaced short palindromic repeat (CRISPR) multilocus sequence typing scheme for subtyping of the major serovars of Salmonella enterica subsp. enterica. Appl Environ Microbiol 77:1946–1956. Shariat N, Kirchner MK, Sandt CH, Trees E, Barrangou R, Dudley EG. 2013. Subtyping of Salmonella enterica serovar Newport outbreak isolates by CRISPR-MVLST and determination of the relationship between CRISPRMVLST and PFGE results. J Clin Microbiol 51:2328– 2336. Liu F, Kariyawasam S, Jayarao BM, Barrangou R, Gerner-Smidt P, Ribot EM, Knabel SJ, Dudley EG. 2011. Subtyping Salmonella enterica serovar Enteritidis isolates from different sources by using sequence typing based on virulence genes and clustered regularly interspaced short palindromic repeats (CRISPRs). Appl Environ Microbiol 77:4520–4526.

10. Molecular Epidemiology n 108. Bennett S. 2004. Solexa Ltd. Pharmacogenomics 5:433– 438. 109. Clarke SC. 2005. Pyrosequencing: nucleotide sequencing technology with bacterial genotyping applications. Expert Rev Mol Diagn 5:947–953. 110. Morozova O, Marra MA. 2008. Applications of nextgeneration sequencing technologies in functional genomics. Genomics 92:255–264. 111. Rothberg JM, Hinz W, Rearick TM, Schultz J, Mileski W, Davey M, Leamon JH, Johnson K, Milgrew MJ, Edwards M, Hoon J, Simons JF, Marran D, Myers JW, Davidson JF, Branting A, Nobile JR, Puc BP, Light D, Clark TA, Huber M, Branciforte JT, Stoner IB, Cawley SE, Lyons M, Fu Y, Homer N, Sedova M, Miao X, Reed B, Sabina J, Feierstein E, Schorn M, Alanjary M, Dimalanta E, Dressman D, Kasinskas R, Sokolsky T, Fidanza JA, Namsaraev E, McKernan KJ, Williams A, Roth GT, Bustillo J. 2011. An integrated semiconductor device enabling non-optical genome sequencing. Nature 475:348–352. 112. Eid J, Fehr A, Gray J, Luong K, Lyle J, Otto G, Peluso P, Rank D, Baybayan P, Bettman B, Bibillo A, Bjornson K, Chaudhuri B, Christians F, Cicero R, Clark S, Dalal R, Dewinter A, Dixon J, Foquet M, Gaertner A, Hardenbol P, Heiner C, Hester K, Holden D, Kearns G, Kong X, Kuse R, Lacroix Y, Lin S, Lundquist P, Ma C, Marks P, Maxham M, Murphy D, Park I, Pham T, Phillips M, Roy J, Sebra R, Shen G, Sorenson J, Tomaney A, Travers K, Trulson M, Vieceli J, Wegener J, Wu D, Yang A, Zaccarin D, Zhao P, Zhong F, Korlach J, Turner S. 2009. Real-time DNA sequencing from single polymerase molecules. Science 323:133–138. 113. Stoddart D, Heron AJ, Mikhailova E, Maglia G, Bayley H. 2009. Single-nucleotide discrimination in immobilized DNA oligonucleotides with a biological nanopore. Proc Natl Acad Sci USA 106:7702–7707. 114. Glenn TC. 2011. Field guide to next-generation DNA sequencers. Mol Ecol Resources 11:759–769. 115. Junemann S, Sedlazeck FJ, Prior K, Albersmeier A, John U, Kalinowski J, Mellmann A, Goesmann A, von Haeseler A, Stoye J, Harmsen D. 2013. Updating benchtop sequencing performance comparison. Nat Biotechnol 31:294–296. 116. Loman NJ, Constantinidou C, Chan JZ, Halachev M, Sergeant M, Penn CW, Robinson ER, Pallen MJ. 2012. High-throughput bacterial genome sequencing: an embarrassment of choice, a world of opportunity. Nat Rev Microbiol 10:599–606. 117. Eyre DW, Golubchik T, Gordon NC, Bowden R, Piazza P, Batty EM, Ip CL, Wilson DJ, Didelot X, O’Connor L, Lay R, Buck D, Kearns AM, Shaw A, Paul J, Wilcox MH, Donnelly PJ, Peto TE, Walker AS, Crook DW. 2012. A pilot study of rapid benchtop sequencing of Staphylococcus aureus and Clostridium difficile for outbreak detection and surveillance. BMJ Open 2(3):pii–e001124. 118. Köser CU, Holden MTG, Ellington MJ, Cartwright EJP, Brown NM, Ogilvy-Stuart AL, Hsu LY, Chewapreecha C, Croucher NJ, Harris SR, Sanders M, Enright MC, Dougan G, Bentley SD, Parkhill J, Fraser LJ, Betley JR, Schulz-Trieglaff OB, Smith GP, Peacock SJ. 2012. Rapid whole-genome sequencing for investigation of a neonatal MRSA outbreak. N Engl J Med 366:2267– 2275. 119. Lewis T, Loman NJ, Bingle L, Jumaa P, Weinstock GM, Mortiboy D, Pallen MJ. 2010. High-throughput whole-genome sequencing to dissect the epidemiology of Acinetobacter baumannii isolates from a hospital outbreak. J Hosp Infect 75:37–41. 120. Millar BC, Xu J, Moore JE. 2007. Molecular diagnostics of medically important bacterial infections. Curr Issues Mol Biol 9:21–39.

155

121. Garten RJ, Davis CT, Russell CA, Shu B, Lindstrom S, Balish A, Sessions WM, Xu X, Skepner E, Deyde V, Okomo-Adhiambo M, Gubareva L, Barnes J, Smith CB, Emery SL, Hillman MJ, Rivailler P, Smagala J, de Graaf M, Burke DF, Fouchier RA, Pappas C, AlpucheAranda CM, Lopez-Gatell H, Olivera H, Lopez I, Myers CA, Faix D, Blair PJ, Yu C, Keene KM, Dotson PD Jr, Boxrud D, Sambol AR, Abid SH, St George K, Bannerman T, Moore AL, Stringer DJ, Blevins P, Demmler-Harrison GJ, Ginsberg M, Kriner P, Waterman S, Smole S, Guevara HF, Belongia EA, Clark PA, Beatrice ST, Donis R, Katz J, Finelli L, Bridges CB, Shaw M, Jernigan DB, Uyeki TM, Smith DJ, Klimov AI, Cox NJ. 2009. Antigenic and genetic characteristics of swineorigin 2009 A(H1N1) influenza viruses circulating in humans. Science 325:197–201. 122. Grad YH, Godfrey P, Cerquiera GC, Mariani-Kurkdjian P, Gouali M, Bingen E, Shea TP, Haas BJ, Griggs A, Young S, Zeng Q, Lipsitch M, Waldor MK, Weill FX, Wortman JR, Hanage WP. 2013. Comparative genomics of recent Shiga toxin-producing Escherichia coli O104:H4: short-term evolution of an emerging pathogen. mBio 4(1):e00452–12. 123. Rasko DA, Webster DR, Sahl JW, Bashir A, Boisen N, Scheutz F, Paxinos EE, Sebra R, Chin CS, Iliopoulos D, Klammer A, Peluso P, Lee L, Kislyuk AO, Bullard J, Kasarskis A, Wang S, Eid J, Rank D, Redman JC, Steyert SR, Frimodt-Moller J, Struve C, Petersen AM, Krogfelt KA, Nataro JP, Schadt EE, Waldor MK. 2011. Origins of the E. coli strain causing an outbreak of hemolytic-uremic syndrome in Germany. N Engl J Med 365:709–717. 124. Gerner-Smidt P, Hise K, Kincaid J, Hunter S, Rolando S, Hyytia-Trees E, Ribot EM, Swaminathan B. 2006. PulseNet USA: a five-year update. Foodborne Pathog Dis 3:9–19. 125. Stucki D, Malla B, Hostettler S, Huna T, Feldmann J, Yeboah-Manu D, Borrell S, Fenner L, Comas I, Coscolla M, Gagneux S. 2012. Two new rapid SNP-typing methods for classifying Mycobacterium tuberculosis complex into the main phylogenetic lineages. PLoS One 7:e41253. 126. van Gent M, Bart MJ, van der Heide HG, Heuvelman KJ, Kallonen T, He Q, Mertsola J, Advani A, Hallander HO, Janssens K, Hermans PW, Mooi FR. 2011. SNPbased typing: a useful tool to study Bordetella pertussis populations. PLoS One 6:e20340. 127. Jolley KA, Maiden MC. 2010. BIGSdb: scalable analysis of bacterial genome variation at the population level. BMC Bioinformatics 11:595. 128. Rasko DA, Myers GS, Ravel J. 2005. Visualization of comparative genomic analyses by BLAST score ratio. BMC Bioinformatics 6:2. 129. Gardner SM, Slezak T. 2010. Scalable SNP analyses of 100+ bacterial or viral genomes. J Forensic Res 1:107. 130. Garaizar J, Rementeria A, Porwollik S. 2006. DNA microarray technology: a new tool for the epidemiological typing of bacterial pathogens? FEMS Immunol Med Microbiol 47:178–189. 131. Dunbar SA. 2006. Applications of Luminex xMAP technology for rapid, high-throughput multiplexed nucleic acid detection. Clin Chim Acta 363:71–82. 132. Borucki MK, Kim SH, Call DR, Smole SC, Pagotto F. 2004. Selective discrimination of Listeria monocytogenes epidemic strains by a mixed-genome DNA microarray compared to discrimination by pulsed-field gel electrophoresis, ribotyping, and multilocus sequence typing. J Clin Microbiol 42:5270–5276. 133. McLoughlin KS. 2011. Microarrays for pathogen detection and analysis. Brief Funct Genomics 10:342–353. 134. Davis MA, Lim JY, Soyer Y, Harbottle H, Chang YF, New D, Orfe LH, Besser TE, Call DR. 2010. Develop-

156 n

135.

136.

137.

138.

139.

140.

141.

142.

143.

144.

145.

146.

147.

148.

149.

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

ment and validation of a resistance and virulence gene microarray targeting Escherichia coli and Salmonella enterica. J Microbiol Methods 82:36–41. Dunbar S, Jacobson J. 2007. Quantitative, multiplexed detection of Salmonella and other pathogens by Luminex xMAP suspension array. Methods Mol Biol 394:1–19. Warsen AE, Krug MJ, LaFrentz S, Stanek DR, Loge FJ, Call DR. 2004. Simultaneous discrimination between 15 fish pathogens by using 16S ribosomal DNA PCR and DNA microarrays. Appl Environ Microbiol 70:4216–4221. Quinones B, Swimley MS, Narm KE, Patel RN, Cooley MB, Mandrell RE. 2012. O-antigen and virulence profiling of Shiga toxin-producing Escherichia coli by a rapid and cost-effective DNA microarray colorimetric method. Front Cell Infect Microbiol 2:61. Borucki MK, Reynolds J, Call DR, Ward TJ, Page B, Kadushin J. 2005. Suspension microarray with dendrimer signal amplification allows direct and high-throughput subtyping of Listeria monocytogenes from genomic DNA. J Clin Microbiol 43:3255–3259. Palacios G, Quan PL, Jabado OJ, Conlan S, Hirschberg DL, Liu Y, Zhai J, Renwick N, Hui J, Hegyi H, Grolla A, Strong JE, Towner JS, Geisbert TW, Jahrling PB, Buchen-Osmond C, Ellerbrok H, Sanchez-Seco MP, Lussier Y, Formenty P, Nichol MS, Feldmann H, Briese T, Lipkin WI. 2007. Panmicrobial oligonucleotide array for diagnosis of infectious diseases. Emerg Infect Dis 13:73– 81. Quan PL, Briese T, Palacios G, Lipkin WI. 2008. Rapid sequence-based diagnosis of viral infection. Antiviral Res 79:1–5. Wang D, Urisman A, Liu YT, Springer M, Ksiazek TG, Erdman DD, Mardis ER, Hickenbotham M, Magrini V, Eldred J, Latreille JP, Wilson RK, Ganem D, DeRisi JL. 2003. Viral discovery and sequence recovery using DNA microarrays. PLoS Biol 1:E2. Brinkman NE, Fout GS. 2009. Development and evaluation of a generic tag array to detect and genotype noroviruses in water. J Virol Methods 156:8–18. Jaaskelainen AJ, Maunula L. 2006. Applicability of microarray technique for the detection of noro- and astroviruses. J Virol Methods 136:210–216. Chiu CY, Alizadeh AA, Rouskin S, Merker JD, Yeh E, Yagi S, Schnurr D, Patterson BK, Ganem D, DeRisi JL. 2007. Diagnosis of a critical respiratory illness caused by human metapneumovirus by use of a pan-virus microarray. J Clin Microbiol 45:2340–2343. Chiu CY, Rouskin S, Koshy A, Urisman A, Fischer K, Yagi S, Schnurr D, Eckburg PB, Tompkins LS, Blackburn BG, Merker JD, Patterson BK, Ganem D, DeRisi JL. 2006. Microarray detection of human parainfluenzavirus 4 infection associated with respiratory failure in an immunocompetent adult. Clin Infect Dis 43:e71–e76. Lin B, Blaney KM, Malanoski AP, Ligler AG, Schnur JM, Metzgar D, Russell KL, Stenger DA. 2007. Using a resequencing microarray as a multiple respiratory pathogen detection assay. J Clin Microbiol 45:443–452. Erlandsson L, Rosenstierne MW, McLoughlin K, Jaing C, Fomsgaard A. 2011. The microbial detection array combined with random Phi29-amplification used as a diagnostic tool for virus detection in clinical samples. PLoS One 6:e22631. Gardner SN, Jaing CJ, McLoughlin KS, Slezak TR. 2010. A microbial detection array (MDA) for viral and bacterial detection. BMC Genomics 11:668. Honma S, Chizhikov V, Santos N, Tatsumi M, Timenetsky Mdo C, Linhares AC, Mascarenhas JD, Ushijima H, Armah GE, Gentsch JR, Hoshino Y. 2007. Development and validation of DNA microarray for genotyping group A rotavirus VP4 (P[4], P[6], P[8], P[9], and P[14])

150.

151.

152.

153.

154.

155.

156.

157.

158.

159.

160.

161.

162.

163.

and VP7 (G1 to G6, G8 to G10, and G12) genes. J Clin Microbiol 45:2641–2648. Huang Y, Tang H, Duffy S, Hong Y, Norman S, Ghosh M, He J, Bose M, Henrickson KJ, Fan J, Kraft AJ, Weisburg WG, Mather EL. 2009. Multiplex assay for simultaneously typing and subtyping influenza viruses by use of an electronic microarray. J Clin Microbiol 47:390– 396. Townsend MB, Dawson ED, Mehlmann M, Smagala JA, Dankbar DM, Moore CL, Smith CB, Cox NJ, Kuchta RD, Rowlen KL. 2006. Experimental evaluation of the FluChip diagnostic microarray for influenza virus surveillance. J Clin Microbiol 44:2863–2871. Pas SD, Tran N, de Man RA, Burghoorn-Maas C, Vernet G, Niesters HG. 2008. Comparison of reverse hybridization, microarray, and sequence analysis for genotyping hepatitis B virus. J Clin Microbiol 46:1268–1273. Sturmer M, Berger A, Preiser W. 2004. HIV-1 genotyping: comparison of two commercially available assays. Expert Rev Mol Diagn 4:281–291. Sergeev N, Rubtcova E, Chizikov V, Schmid DS, Loparev VN. 2006. New mosaic subgenotype of varicellazoster virus in the USA: VZV detection and genotyping by oligonucleotide-microarray. J Virol Methods 136:8–16. Neverov AA, Riddell MA, Moss WJ, Volokhov DV, Rota PA, Lowe LE, Chibo D, Smit SB, Griffin DE, Chumakov KM, Chizhikov VE. 2006. Genotyping of measles virus in clinical specimens on the basis of oligonucleotide microarray hybridization patterns. J Clin Microbiol 44:3752–3759. Feng Q, Cherne S, Winer RL, Balasubramanian A, Lee SK, Hawes SE, Kiviat NB, Koutsky LA. 2009. Development and evaluation of a liquid bead microarray assay for genotyping genital human papillomaviruses. J Clin Microbiol 47:547–553. Lee JK, Kim MK, Song SH, Hong JH, Min KJ, Kim JH, Song ES, Lee J, Lee JM, Hur SY. 2009. Comparison of human papillomavirus detection and typing by hybrid capture 2, linear array, DNA chip, and cycle sequencing in cervical swab samples. Int J Gynecol Cancer 19:266– 272. Cherkasova E, Laassri M, Chizhikov V, Korotkova E, Dragunsky E, Agol VI, Chumakov K. 2003. Microarray analysis of evolution of RNA viruses: evidence of circulation of virulent highly divergent vaccine-derived polioviruses. Proc Natl Acad Sci USA 100:9398–9403. Fitzgerald C, Collins M, van Duyne S, Mikoleit M, Brown T, Fields P. 2007. Multiplex, bead-based suspension array for molecular determination of common Salmonella serogroups. J Clin Microbiol 45:3323–3334. Jackson SA, Patel IR, Barnaba T, LeClerc JE, Cebula TA. 2011. Investigating the global genomic diversity of Escherichia coli using a multi-genome DNA microarray platform with novel gene prediction strategies. BMC Genomics 12:349. Laksanalamai P, Jackson SA, Mammel MK, Datta AR. 2012. High density microarray analysis reveals new insights into genetic footprints of Listeria monocytogenes strains involved in listeriosis outbreaks. PLoS One 7:e32896. Scaria J, Palaniappan RU, Chiu D, Phan JA, Ponnala L, McDonough P, Grohn YT, Porwollik S, McClelland M, Chiou CS, Chu C, Chang YF. 2008. Microarray for molecular typing of Salmonella enterica serovars. Mol Cell Probes 22:238–243. Frye JG, Lindsey RL, Rondeau G, Porwollik S, Long F, McClelland M, Jackson CR, Englen MD, Meinersmann RJ, Berrang ME, Davis JA, Barrett JB, Turpin JB, Thitaram SN, Fedorka-Cray PJ. 2010. Development of a DNA microarray to detect antimicrobial resistance

10. Molecular Epidemiology n

164.

165.

166.

167.

168.

169.

170.

171.

172.

173.

174.

175.

176.

genes identified in the National Center for Biotechnology Information database. Microb Drug Resist 16:9–19. Lascols C, Hackel M, Hujer AM, Marshall SH, Bouchillon SK, Hoban DJ, Hawser SP, Badal RE, Bonomo RA. 2012. Using nucleic acid microarrays to perform molecular epidemiology and detect novel beta-lactamases: a snapshot of extended-spectrum beta-lactamases throughout the world. J Clin Microbiol 50:1632–1639. Peter H, Berggrav K, Thomas P, Pfeifer Y, Witte W, Templeton K, Bachmann TT. 2012. Direct detection and genotyping of Klebsiella pneumoniae carbapenemases from urine by use of a new DNA microarray test. J Clin Microbiol 50:3990–3998. Zimenkov DV, Antonova OV, Kuz’min AV, Isaeva YD, Krylova LY, Popov SA, Zasedatelev AS, Mikhailovich VM, Gryadunov DA. 2013. Detection of second-line drug resistance in Mycobacterium tuberculosis using oligonucleotide microarrays. BMC Infect Dis 13:240. Sulaiman IM, Liu X, Frace M, Sulaiman N, OlsenRasmussen M, Neuhaus E, Rota PA, Wohlhueter RM. 2006. Evaluation of affymetrix severe acute respiratory syndrome resequencing GeneChips in characterization of the genomes of two strains of coronavirus infecting humans. Appl Environ Microbiol 72:207–211. Sulaiman IM, Sammons SA, Wohlhueter RM. 2008. Smallpox virus resequencing GeneChips can also rapidly ascertain species status for some zoonotic non-variola orthopoxviruses. J Clin Microbiol 46:1507–1509. Dotsch A, Pommerenke C, Bredenbruch F, Geffers R, Haussler S. 2009. Evaluation of a microarray-hybridization based method applicable for discovery of single nucleotide polymorphisms (SNPs) in the Pseudomonas aeruginosa genome. BMC Genomics 10:29. Zhang W, Qi W, Albert TJ, Motiwala AS, Alland D, Hyytia-Trees EK, Ribot EM, Fields PI, Whittam TS, Swaminathan B. 2006. Probing genomic diversity and evolution of Escherichia coli O157 by single nucleotide polymorphisms. Genome Res 16:757–767. Westblade LF, Jennemann R, Branda JA, Bythrow M, Ferraro MJ, Garner OB, Ginocchio CC, Lewinski MA, Manji R, Mochon AB, Procop GW, Richter SS, Rychert JA, Sercia L, Burnham CA. 2013. Multicenter study evaluating the Vitek MS system for identification of medically important yeasts. J Clin Microbiol 51:2267–2272. Berrazeg M, Diene SM, Drissi M, Kempf M, Richet H, Landraud L, Rolain JM. 2013. Biotyping of multidrugresistant Klebsiella pneumoniae clinical isolates from France and Algeria using MALDI-TOF MS. PLoS One 8:e61428. Clark CG, Kruczkiewicz P, Guan C, McCorrister SJ, Chong P, Wylie J, van Caeseele P, Tabor HA, Snarr P, Gilmour MW, Taboada EN, Westmacott GR. 2013. Evaluation of MALDI-TOF mass spectroscopy methods for determination of Escherichia coli pathotypes. J Microbiol Methods 94:180–191. Mencacci A, Monari C, Leli C, Merlini L, De Carolis E, Vella A, Cacioni M, Buzi S, Nardelli E, Bistoni F, Sanguinetti M, Vecchiarelli A. 2013. Typing of nosocomial outbreaks of Acinetobacter baumannii by use of matrixassisted laser desorption ionization–time of flight mass spectrometry. J Clin Microbiol 51:603–606. Tamura H, Hotta Y, Sato H. 2013. Novel accurate bacterial discrimination by MALDI-time-of-flight MS based on ribosomal proteins coding in S10-spc-alpha operon at strain level S10-GERMS. J Am Soc Mass Spectrom 24:1185–1193. Jacob D, Sauer U, Housley R, Washington C, SannesLowery K, Ecker DJ, Sampath R, Grunow R. 2012. Rapid and high-throughput detection of highly pathogenic bacteria by Ibis PLEX-ID technology. PLoS One 7:e39928.

157

177. Richmond GS, Khine H, Zhou TT, Ryan DE, Brand T, McBride MT, Killeen K. 2011. MassCode liquid arrays as a tool for multiplexed high-throughput genetic profiling. PLoS One 6:e18967. 178. Syrmis MW, Moser RJ, Whiley DM, Vaska V, Coombs GW, Nissen MD, Sloots TP, Nimmo GR. 2011. Comparison of a multiplexed MassARRAY system with realtime allele-specific PCR technology for genotyping of methicillin-resistant Staphylococcus aureus. Clin Microbiol Infect 17:1804–1810. 179. Grimont PAD, Weill FX. 2007. Antigenic Formulae of the Salmonella Serovars, 9th ed. WHO Collaborating Centre for Reference and Research on Salmonella, Institut Pasteur, Paris, France. 180. Ranieri ML, Shi C, Moreno Switt AI, den Bakker HC, Wiedmann M. 2013. Comparison of typing methods with a new procedure based on sequence characterization for Salmonella serovar prediction. J Clin Microbiol 51:1786– 1797. 181. McQuiston JR, Waters RJ, Dinsmore BA, Mikoleit ML, Fields PI. 2011. Molecular determination of H antigens of Salmonella by use of a microsphere-based liquid array. J Clin Microbiol 49:565–573. 182. Seong WJ, Kwon HJ, Kim TE, Lee DY, Park MS, Kim JH. 2012. Molecular serotyping of Salmonella enterica by complete rpoB gene sequencing. J Microbiol 50:962–969. 183. Gerner-Smidt P, Kincaid J, Kubota K, Hise K, Hunter SB, Fair MA, Norton D, Woo-Ming A, Kurzynski T, Sotir MJ, Head M, Holt K, Swaminathan B. 2005. Molecular surveillance of Shiga toxigenic Escherichia coli O157 by PulseNet USA. J Food Prot 68:1926–1931. 184. Tenover FC, Arbeit RD, Goering RV, Mickelsen PA, Murray BE, Persing DH, Swaminathan B. 1995. Interpreting chromosomal DNA restriction patterns produced by pulsed-field gel electrophoresis: criteria for bacterial strain typing. J Clin Microbiol 33:2233–2239. 185. Donnarumma F, Indorato C, Mastromei G, Goti E, Nicoletti P, Pecile P, Fanci R, Bosi A, Casalone E. 2012. Molecular analysis of population structure and antibiotic resistance of Klebsiella isolates from a three-year surveillance program in Florence hospitals, Italy. Eur J Clin Microbiol Infect Dis 31:371–378. 186. Fontana C, Favaro M, Minelli S, Bossa MC, Testore GP, Leonardis F, Natoli S, Favalli C. 2008. Acinetobacter baumannii in intensive care unit: a novel system to study clonal relationship among the isolates. BMC Infect Dis 8:79. 187. Vigil KJ, Adachi JA, Aboufaycal H, Hachem RY, Reitzel RA, Jiang Y, Tarrand JJ, Chemaly RF, Bodey GP, Rolston KV, Raad I. 2009. Multidrug-resistant Escherichia coli bacteremia in cancer patients. Am J Infect Control 30:30. 188. Patel MM, Hall AJ, Vinje J, Parashar UD. 2009. Noroviruses: a comprehensive review. J Clin Virol 44:1–8. 189. Dowell SF, Groves C, Kirkland KB, Cicirello HG, Ando T, Jin Q, Gentsch JR, Monroe SS, Humphrey CD, Slemp C, Dwyer DM, Meriwether RA, Glass RI. 1995. A multistate outbreak of oyster-associated gastroenteritis: implications for interstate tracing of contaminated shellfish. J Infect Dis 171:1497–1503. 190. Parashar UD, Dow L, Fankhauser RL, Humphrey CD, Miller J, Ando T, Williams KS, Eddy CR, Noel JS, Ingram T, Bresee JS, Monroe SS, Glass RI. 1998. An outbreak of viral gastroenteritis associated with consumption of sandwiches: implications for the control of transmission by food handlers. Epidemiol Infect 121:615–621. 191. Rota PA, Featherstone DA, Bellini WJ. 2009. Molecular epidemiology of measles virus. Curr Top Microbiol Immunol 330:129–150. 192. Pires SM, Evers EG, van Pelt W, Ayers T, Scallan E, Angulo FJ, Havelaar A, Hald T. 2009. Attributing the

158 n

193.

194.

195.

196.

197.

198.

199.

200.

201.

202.

203.

204. 205.

206.

207.

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

human disease burden of foodborne infections to specific sources. Foodborne Pathog Dis 6:417–424. Sheppard SK, Dallas JF, Strachan NJ, MacRae M, McCarthy ND, Wilson DJ, Gormley FJ, Falush D, Ogden ID, Maiden MC, Forbes KJ. 2009. Campylobacter genotyping to determine the source of human infection. Clin Infect Dis 48:1072–1078. Muellner P, Pleydell E, Pirie R, Baker MG, Campbell D, Carter PE, French NP. 2013. Molecular-based surveillance of campylobacteriosis in New Zealand—from source attribution to genomic epidemiology. Euro Surveill 18(3):pii-20365. Mughini Gras L, Smid JH, Wagenaar JA, de Boer AG, Havelaar AH, Friesema IH, French NP, Busani L, van Pelt W. 2012. Risk factors for campylobacteriosis of chicken, ruminant, and environmental origin: a combined case-control and source attribution analysis. PLoS One 7:e42599. Drummond AJ, Rambaut A. 2007. BEAST: Bayesian evolutionary analysis by sampling trees. BMC Evol Biol 7:214. Grenfell BT, Pybus OG, Gog JR, Wood JL, Daly JM, Mumford JA, Holmes EC. 2004. Unifying the epidemiological and evolutionary dynamics of pathogens. Science 303:327–332. Pybus OG, Drummond AJ, Nakano T, Robertson BH, Rambaut A. 2003. The epidemiology and iatrogenic transmission of hepatitis C virus in Egypt: a Bayesian coalescent approach. Mol Biol Evol 20:381–387. Lewis F, Hughes GJ, Rambaut A, Pozniak A, Leigh Brown AJ. 2008. Episodic sexual transmission of HIV revealed by molecular phylodynamics. PLoS Med 5:e50. Smith GJ, Vijaykrishna D, Bahl J, Lycett SJ, Worobey M, Pybus OG, Ma SK, Cheung CL, Raghwani J, Bhatt S, Peiris JS, Guan Y, Rambaut A. 2009. Origins and evolutionary genomics of the 2009 swine-origin H1N1 influenza A epidemic. Nature 459:1122–1125. Lanciotti RS, Ebel GD, Deubel V, Kerst AJ, Murri S, Meyer R, Bowen M, McKinney N, Morrill WE, Crabtree MB, Kramer LD, Roehrig JT. 2002. Complete genome sequences and phylogenetic analysis of West Nile virus strains isolated from the United States, Europe, and the Middle East. Virology 298:96–105. Roehrig JT, Layton M, Smith P, Campbell GL, Nasci R, Lanciotti RS. 2002. The emergence of West Nile virus in North America: ecology, epidemiology, and surveillance. Curr Top Microbiol Immunol 267:223–240. Lal RB, Chakrabarti S, Yang C. 2005. Impact of genetic diversity of HIV-1 on diagnosis, antiretroviral therapy & vaccine development. Indian J Med Res 121:287–314. Bellini WJ, Rota PA. 2011. Biological feasibility of measles eradication. Virus Res 162:72–79. Rota PA, Brown K, Mankertz A, Santibanez S, Shulga S, Muller CP, Hübschen JM, Siqueira M, Beirnes J, Ahmed H, Triki H, Al-Busaidy S, Dosseh A, Byabamazima C, Smit S, Akoua-Koffi C, Bwogi J, Bukenya H, Wairagkar N, Ramamurty N, Incomserb P, Pattamadilok S, Jee Y, Lim W, Xu W, Komase K, Takeda M, Tran T, Castillo-Solorzano C, Chenoweth P, Brown D, Mulders MN, Bellini WJ, Featherstone D. 2011. Global distribution of measles genotypes and measles molecular epidemiology. J Infect Dis 204:S514–S523. Rota PA, Brown KE, Hübschen JM, Muller CP, Icenogle J, Chen M-H, Bankamp B, Kessler JR, Brown DW, Bellini WJ, Featherstone D. 2011. Improving global virologic surveillance for measles and rubella. J Infect Dis 204:S506–S513. Holmes EC, Rambaut A. 2004. Viral evolution and the emergence of SARS coronavirus. Philos Trans R Soc Lond B Biol Sci 359:1059–1065.

208. Ksiazek TG, Erdman D, Goldsmith CS, Zaki SR, Peret T, Emery S, Tong S, Urbani C, Comer JA, Lim W, Rollin PE, Dowell SF, Ling AE, Humphrey CD, Shieh WJ, Guarner J, Paddock CD, Rota P, Fields B, DeRisi J, Yang JY, Cox N, Hughes JM, LeDuc JW, Bellini WJ, Anderson LJ. 2003. A novel coronavirus associated with severe acute respiratory syndrome. N Engl J Med 348:1953–1966. 209. Rota PA, Oberste MS, Monroe SS, Nix WA, Campagnoli R, Icenogle JP, Penaranda S, Bankamp B, Maher K, Chen MH, Tong S, Tamin A, Lowe L, Frace M, DeRisi JL, Chen Q, Wang D, Erdman DD, Peret TC, Burns C, Ksiazek TG, Rollin PE, Sanchez A, Liffick S, Holloway B, Limor J, McCaustland K, Olsen-Rasmussen M, Fouchier R, Gunther S, Osterhaus AD, Drosten C, Pallansch MA, Anderson LJ, Bellini WJ. 2003. Characterization of a novel coronavirus associated with severe acute respiratory syndrome. Science 300:1394–1399. 210. Corman VM, Eckerle I, Bleicker T, Zaki A, Landt O, Eschbach-Bludau M, van Boheemen S, Gopal R, Ballhause M, Bestebroer TM, Muth D, Muller MA, Drexler JF, Zambon M, Osterhaus AD, Fouchier RM, Drosten C. 2012. Detection of a novel human coronavirus by realtime reverse-transcription polymerase chain reaction. Euro Surveill 17(39):pii–20285. 211. van Boheemen S, de Graaf M, Lauber C, Bestebroer TM, Raj VS, Zaki AM, Osterhaus AD, Haagmans BL, Gorbalenya AE, Snijder EJ, Fouchier RA. 2012. Genomic characterization of a newly discovered coronavirus associated with acute respiratory distress syndrome in humans. mBio 3(6):e00473–12. 212. Zaki AM, van Boheemen S, Bestebroer TM, Osterhaus AD, Fouchier RA. 2012. Isolation of a novel coronavirus from a man with pneumonia in Saudi Arabia. N Engl J Med 367:1814–1820. 213. Galea SA, Sweet A, Beninger P, Steinberg SP, Larussa PS, Gershon AA, Sharrar RG. 2008. The safety profile of varicella vaccine: a 10-year review. J Infect Dis 197(Suppl 2):S165–S169. 214. Minor P. 2009. Vaccine-derived poliovirus (VDPV): impact on poliomyelitis eradication. Vaccine 27:2649–2652. 215. Yang CF, Chen HY, Jorba J, Sun HC, Yang SJ, Lee HC, Huang YC, Lin TY, Chen PJ, Shimizu H, Nishimura Y, Utama A, Pallansch M, Miyamura T, Kew O, Yang JY. 2005. Intratypic recombination among lineages of type 1 vaccine-derived poliovirus emerging during chronic infection of an immunodeficient patient. J Virol 79:12623–12634. 216. Arita M, Zhu SL, Yoshida H, Yoneyama T, Miyamura T, Shimizu H. 2005. A Sabin 3-derived poliovirus recombinant contained a sequence homologous with indigenous human enterovirus species C in the viral polymerase coding region. J Virol 79:12650–12657. 217. Belongia EA, Kieke BA, Donahue JG, Greenlee RT, Balish A, Foust A, Lindstrom S, Shay DK. 2009. Effectiveness of inactivated influenza vaccines varied substantially with antigenic match from the 2004-2005 season to the 2006-2007 season. J Infect Dis 199:159–167. 218. Patel M, Pedreira C, De Oliveira LH, Tate J, Orozco M, Mercado J, Gonzalez A, Malespin O, Amador JJ, Umana J, Balmaseda A, Perez MC, Gentsch J, Kerin T, Hull J, Mijatovic S, Andrus J, Parashar U. 2009. Association between pentavalent rotavirus vaccine and severe rotavirus diarrhea among children in Nicaragua. JAMA 301:2243–2251. 219. Patel MM, Clark AD, Glass RI, Greenberg H, Tate J, Santosham M, Sanderson CF, Steele D, Cortese M, Parashar UD. 2009. Broadening the age restriction for initiating rotavirus vaccination in regions with high ro-

10. Molecular Epidemiology n

220.

221.

222.

223.

224.

225.

226.

227.

228.

229.

230.

231.

232.

tavirus mortality: benefits of mortality reduction versus risk of fatal intussusception. Vaccine 27:2916–2922. Metzker ML, Mindell DP, Liu XM, Ptak RG, Gibbs RA, Hillis DM. 2002. Molecular evidence of HIV-1 transmission in a criminal case. Proc Natl Acad Sci USA 99:14292–14297. Lemey P, Van Dooren S, Van Laethem K, Schrooten Y, Derdelinckx I, Goubau P, Brun-Vezinet F, Vaira D, Vandamme AM. 2005. Molecular testing of multiple HIV-1 transmissions in a criminal case. AIDS 19:1649– 1658. Keim P, Van Ert MN, Pearson T, Vogler AJ, Huynh LY, Wagner DM. 2004. Anthrax molecular epidemiology and forensics: using the appropriate marker for different evolutionary scales. Infect Genet Evol 4:205–213. Török TJ, Tauxe RV, Wise RP, Livengood JR, Sokolow R, Mauvais S, Birkness KA, Skeels MR, Horan JM, Foster LR. 1997. A large community outbreak of salmonellosis caused by intentional contamination of restaurant salad bars. JAMA 278:389–395. Achtman M, Zurth K, Morelli G, Torrea G, Guiyoule A, Carniel E. 1999. Yersinia pestis, the cause of plague, is a recently emerged clone of Yersinia pseudotuberculosis. Proc Natl Acad Sci USA 96:14043–14048. Bono JL, Smith TP, Keen JE, Harhay GP, McDaneld TG, Mandrell RE, Jung WK, Besser TE, Gerner-Smidt P, Bielaszewska M, Karch H, Clawson ML. 2012. Phylogeny of Shiga toxin-producing Escherichia coli O157 isolated from cattle and clinically ill humans. Mol Biol Evol 29:2047–2062. Eppinger M, Mammel MK, Leclerc JE, Ravel J, Cebula TA. 2011. Genome signatures of Escherichia coli O157:H7 isolates from the bovine host reservoir. Appl Environ Microbiol 77:2916–2925. Leopold SR, Magrini V, Holt NJ, Shaikh N, Mardis ER, Cagno J, Ogura Y, Iguchi A, Hayashi T, Mellmann A, Karch H, Besser TE, Sawyer SA, Whittam TS, Tarr PI. 2009. A precise reconstruction of the emergence and constrained radiations of Escherichia coli O157 portrayed by backbone concatenomic analysis. Proc Natl Acad Sci USA 106:8713–8718. Holt KE, Parkhill J, Mazzoni CJ, Roumagnac P, Weill FX, Goodhead I, Rance R, Baker S, Maskell DJ, Wain J, Dolecek C, Achtman M, Dougan G. 2008. Highthroughput sequencing provides insights into genome variation and evolution in Salmonella Typhi. Nat Genet 40:987–993. Pearson T, Busch JD, Ravel J, Read TD, Rhoton SD, U’Ren JM, Simonson TS, Kachur SM, Leadem RR, Cardon ML, Van Ert MN, Huynh LY, Fraser CM, Keim P. 2004. Phylogenetic discovery bias in Bacillus anthracis using single-nucleotide polymorphisms from whole-genome sequencing. Proc Natl Acad Sci USA 101:13536– 13541. Kenefic LJ, Beaudry J, Trim C, Daly R, Parmar R, Zanecki S, Huynh L, Van Ert MN, Wagner DM, Graham T, Keim P. 2008. High resolution genotyping of Bacillus anthracis outbreak strains using four highly mutable single nucleotide repeat markers. Lett Appl Microbiol 46:600–603. Vogler AJ, Chan F, Wagner DM, Roumagnac P, Lee J, Nera R, Eppinger M, Ravel J, Rahalison L, Rasoamanana BW, Beckstrom-Sternberg SM, Achtman M, Chanteau S, Keim P. 2011. Phylogeography and molecular epidemiology of Yersinia pestis in Madagascar. PLoS Negl Trop Dis 5:e1319. Hendriksen RS, Hyytia-Trees E, Pulsrikarn C, Pornruangwong S, Chaichana P, Svendsen CA, Ahmed R, Mikoleit M. 2012. Characterization of Salmonella enterica serovar Enteritidis isolates recovered from blood and stool specimens in Thailand. BMC Microbiol 12:92.

159

233. Barrett TJ, Gerner-Smidt P, Swaminathan B. 2006. Interpretation of pulsed-field gel electrophoresis patterns in foodborne disease investigations and surveillance. Foodborne Pathog Dis 3:20–31. 234. de Valk HA, Meis JF, Bretagne S, Costa JM, Lasker BA, Balajee SA, Pasqualotto AC, Anderson MJ, Alcazar-Fuoli L, Mellado E, Klaassen CH. 2009. Interlaboratory reproducibility of a microsatellite-based typing assay for Aspergillus fumigatus through the use of allelic ladders: proof of concept. Clin Microbiol Infect 15:180–187. 235. Aanensen DM, Spratt BG. 2005. The multilocus sequence typing network: mlst.net. Nucleic Acids Res 33:W728–W733. 236. Mavromatis K, Land ML, Brettin TS, Quest DJ, Copeland A, Clum A, Goodwin L, Woyke T, Lapidus A, Klenk HP, Cottingham RW, Kyrpides NC. 2012. The fast changing landscape of sequencing technologies and their impact on microbial genome assemblies and annotation. PLoS One 7:e48837. 237. Allard MW, Luo Y, Strain E, Pettengill J, Timme R, Wang C, Li C, Keys CE, Zheng J, Stones R, Wilson MR, Musser SM, Brown EW. 2013. On the evolutionary history, population genetics and diversity among isolates of Salmonella Enteritidis PFGE pattern JEGX01.0004. PLoS One 8:e55254. 238. Bielaszewska M, Prager R, Zhang W, Friedrich AW, Mellmann A, Tschape H, Karch H. 2006. Chromosomal dynamism in progeny of outbreak-related sorbitol-fermenting enterohemorrhagic Escherichia coli O157:NM. Appl Environ Microbiol 72:1900–1909. 239. Jolley KA, Chan MS, Maiden MC. 2004. mlstdbNetdistributed multi-locus sequence typing (MLST) databases. BMC Bioinformatics 5:86. 240. Friedrich AW, Witte W, de Lencastre H, Hryniewicz W, Scheres J, Westh H. 2008. A European laboratory network for sequence-based typing of methicillin-resistant Staphylococcus aureus (MRSA) as a communication platform between human and veterinary medicine—an update on SeqNet.org. Euro Surveill 13(19):pii–18862. 241. Oliveira DC, Milheirico C, Vinga S, de Lencastre H. 2006. Assessment of allelic variation in the ccrAB locus in methicillin-resistant Staphylococcus aureus clones. J Antimicrob Chemother 58:23–30. 242. Struelens MJ. 1996. Consensus guidelines for appropriate use and evaluation of microbial epidemiologic typing systems. Clin Microbiol Infect 2:2–11. 243. Reddy TB, Riley R, Wymore F, Montgomery P, DeCaprio D, Engels R, Gellesch M, Hubble J, Jen D, Jin H, Koehrsen M, Larson L, Mao M, Nitzberg M, Sisk P, Stolte C, Weiner B, White J, Zachariah ZK, Sherlock G, Galagan JE, Ball CA, Schoolnik GK. 2009. TB database: an integrated platform for tuberculosis research. Nucleic Acids Res 37:D499–D508. 244. Swaminathan B, Gerner-Smidt P, Ng LK, Lukinmaa S, Kam KM, Rolando S, Gutierrez EP, Binsztein N. 2006. Building PulseNet International: an interconnected system of laboratory networks to facilitate timely public health recognition and response to foodborne disease outbreaks and emerging foodborne diseases. Foodborne Pathog Dis 3:36–50. 245. CDC. 2003. Norovirus activity—United States, 2002. MMWR Morb Mortal Wkly Rep 52:1–5. 246. Koopmans M, Vennema H, Heersma H, van Strien E, van Duynhoven Y, Brown D, Reacher M, Lopman B. 2003. Early identification of common-source foodborne virus outbreaks in Europe. Emerg Infect Dis 9:1136–1142. 247. Li Y, Gao L, Ding Y, Xu Y, Zhou M, Huang W, Jing Y, Li H, Wang L, Yu L. 2013. Establishment and application of real-time quantitative PCR for diagnosing invasive Aspergillosis via the blood in hematological patients: tar-

160 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

geting a specific sequence of Aspergillus 28S-ITS2. BMC Infect Dis 13:255. 248. van der Helm JJ, Sabajo LO, Grunberg AW, Morre SA, Speksnijder AG, de Vries HJ. 2012. Point-of-care test for detection of urogenital chlamydia in women shows low sensitivity. A performance evaluation study in two clinics in Suriname. PLoS One 7:e32122. 249. Yakoob J, Abbas Z, Khan R, Naz S, Ahmad Z, Islam M, Awan S, Jafri F, Jafri W. 2012. Prevalence of non Helicobacter pylori species in patients presenting with dyspepsia. BMC Gastroenterol 12:3. 250. Chen EC, Miller SA, DeRisi JL, Chiu CY. 2011. Using a pan-viral microarray assay (Virochip) to screen clinical samples for viral pathogens. J Vis Exp 50:e2536

251. Claas EC, Burnham C-AD, Mazzulli T, Templeton K, Topin F. 2013. Performance of the xTAG® gastrointestinal pathogen panel, a multiplex molecular assay for simultaneous detection of bacterial, viral, and parasitic causes of infectious gastroenteritis. J Microbiol Biotechnol 23:1041. 252. Bertelli C, Greub G. 2013. Rapid bacterial genome sequencing: methods and applications in clinical microbiology. Clin Microbiol Infect 19:803–813. 253. Lagier JC, Armougom F, Million M, Hugon P, Pagnier I, Robert C, Bittar F, Fournous G, Gimenez G, Maraninchi M, Trape JF, Koonin EV, La Scola B, Raoult D. 2012. Microbial culturomics: paradigm shift in the human gut microbiome study. Clin Microbiol Infect 18:1185– 1193.

Procedures for the Storage of Microorganisms* ROSEMARY C. SHE AND CATHY A. PETTI

11 Long- and short-term preservation of microorganisms for future study has a long tradition in microbiology. As early as the 1880s, it was observed that bacteria survived well in ice and that freeze-thaw cycles damaged the cell wall (1). By the 1900s, researchers had optimized methods to preserve tissues, viruses, and bacteria for later use by freezing and drying. It had become apparent to early researchers that preserving microorganisms allowed them to be studied at a later time and to remain viable when transported to others (2). Indeed, culture collections of microorganisms are valuable resources for scientific research in microbial diversity and evolution, patient care management, epidemiological investigations, and educational purposes. Preserved individual strains of microorganisms serve as permanent records of microorganisms’ unique phenotypic profiles and provide the material for further genotypic characterizations. Such reference collections can encompass rare infectious agents unique to an individual or catalog the history of disease caused by common pathogens such as those responsible for community outbreaks. There are multiple methods for microbial preservation. Effective storage is defined by the ability to maintain an organism in a viable state free of contamination and without changes in its genotypic or phenotypic characteristics. Secondly, the organism must be easily restored to its condition prior to preservation. Microbial preservation methods have been evaluated extensively over the past 60 years, and often, optimal methods for preservation depend on a microorganism’s taxonomic classification. Review articles, monographs, and books have been published that provide detailed information about the storage of various types of microorganisms (3–7). For clinical microbiology laboratories, simple and broadly applied methods are necessary to maintain organisms for short- and long-term recovery. This chapter presents methods that can be used for the storage of bacteria, protozoa, fungi, and viruses.

odic subculture to fresh medium. Although simple, if microorganisms are saved for >1 week, this method is potentially labor-intensive, requires extensive laboratory space, and may compromise a microorganism’s phenotypic profile. Each transfer to a new subculture increases the likelihood of mutation with undesirable changes in a microorganism’s characteristics. Furthermore, plasmids may be lost with subculturing. The interval between transfers varies among organisms. Additionally, the rate of mutation is quite variable. Some organisms appear stable indefinitely with repeated transfer, and others may change phenotypic traits after as few as two or three passages. The actual rate of mutation, however, has not been studied until recently using sequencing technology (8, 9). Issues that must be addressed with direct transfer include the medium to be used, the storage conditions, and the frequency of transfer.

Maintenance Medium The medium should support the survival of the microorganism but minimize its metabolic processes and slow its rate of growth. Extreme environments should be avoided because microorganisms have the unique ability to adapt through mutation events in order to survive in suboptimal surroundings. A medium with too high a nutrient content will induce rapid replication that requires more frequent transfers. The optimal medium for maintaining microorganisms has not been clearly defined and most likely varies from one species to another and may even depend on the individual strain. Media that have been used include distilled water, tryptic soy broth, and nutrient broths, all of which may be used with or without cryopreservatives.

Storage Conditions Many laboratories store organisms, most often bacteria, for short periods on routine agar media at the workbench. Cultures kept in this fashion are subject to drying. A better method is to transfer organisms into an agar slant tube with a screw top and to store them in an organized location away from light and significant temperature changes. To prevent drying, caps can include rubber liners, or film can be wrapped over the top of the tube before or after the cap is screwed on. Storage at lower temperatures (5 to 8°C) slows metabolic processes and maintains viability for longer periods.

OVERVIEW OF PRESERVATION METHODS Short-Term Preservation Methods Direct Transfer to Subculture The simplest method for maintaining the short-term viability of microorganisms, most often used for bacteria, is peri-

Frequency of Transfer There is no set protocol for the frequency of transfer since storage conditions, media used, and types of microorganisms

*This chapter contains information presented by Cathy A. Petti and Karen C. Carroll in chapter 9 of the 10th edition of this Manual.

doi:10.1128/9781555817381.ch11

161

162 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

vary among laboratories. Individual laboratories may conduct studies for each category of microorganism to determine optimal intervals between transfers under their conditions used for storage. Such studies would involve performing subcultures at scheduled times until the laboratory identifies an acceptable interval between transfers at which a microorganism can reliably and reproducibly be recovered. (When transfers are performed, 5 to 10 representative colonies should be used to avoid the possibility of introducing an altered genotypic or phenotypic characteristic.) For the passage and storage of quality control organisms for antimicrobial susceptibility testing, published standards should be followed (10).

Quality Control Procedures Although it is not necessary with each transfer, the status of the specimen should be assessed periodically. Ongoing viability, stability of phenotype, microorganism identity, and the rate of contamination of specimens should be determined and noted in a log.

long periods (4). Instead, commercial silica gel can be used in small cotton-plugged tubes after being heated in an oven to 175°C for 1.5 to 2 h (6), with moderately successful recovery of fungi. Alternatively, a suspension of 108 microorganisms can be inoculated onto sterile filter paper strips or disks. The paper is dried in air or under a vacuum and is placed in sterile vials. These vials can be stored in the refrigerator for up to 4 years, and then single strips or disks can be removed as needed (4). This method is commonly used for quality control organisms.

Storage in Distilled Water Most organisms do poorly in distilled water, but some survive for prolonged periods. Many fungi and Pseudomonas spp. survive for several years in distilled water at room temperature (6, 13). McGinnis et al. found that with the exception of fungi that do not easily sporulate, 93% of yeasts, molds, and aerobic actinomycetes can be easily and inexpensively preserved this way (14).

Long-Term Preservation Methods Immersion in Oil An alternative to capping tubes is to add a layer of mineral oil to the top of the specimen. Many bacteria and fungi can be stored for periods of up to 2 to 3 years by this method, and transfers are not needed as frequently. Microorganisms are still metabolically active in this environment, and mutations can still occur. Mineral oil should be medicinal-grade oil with a specific gravity of 0.865 to 0.890. Contamination of the specimen can occur if the mineral oil is not adequately sterilized. For sterilization, it should be heated to 170°C for 1 to 2 h in an oven (4). Autoclaving is not considered acceptable. Sterile mineral oil is also commercially available. To prepare the specimen, an inoculum of 5 to 10 colonies of the microorganism should be placed on an agar slant or in tubed broth media. Once growth is identified, a layer of mineral oil at least 1 to 2 cm deep is added, and the agar must not be exposed to air. As with the simple transfer method, tests for viability should be performed to determine the optimal transfer schedule that will ensure microorganism recovery. Transfers will be less frequent than those of microorganisms stored without oil; however, oil is more difficult to add to vials and to clean up in the event of spills.

Freezing at −20°C Refrigeration or freezing in ordinary freezers at −20°C may be used to preserve microorganisms for periods longer than those that can be accomplished by repeated transfers. Viability may be maintained for as long as 1 to 2 years for specific microorganisms, but overall, damage caused by ice crystal formation (6) and electrolyte fluctuations (4) results in poor long-term survival. The medium used for storage appears to be important, since preservation times vary from a few months to 2 years depending upon which medium is used (6, 11, 12). Modern self-defrosting freezers with freeze-thaw cycles must be avoided because cyclic temperature fluctuation will destroy the microorganism.

Drying Although most microorganisms do not survive drying, molds and some spore-forming bacteria may be dried and stored for prolonged periods. Soil has been described as a storage medium if it is autoclaved and air dried, but it is not a standardized, defined, and consistent product for use over

Whereas the methods described above may be used to store microorganisms for periods of up to a few years, ultralowtemperature freezing and freeze-drying (lyophilization) are recommended for long-term storage. Although the initial investment in ultralow-temperature freezers and lyophilization may be costly, these methods are less labor-intensive over time, require less laboratory space (e.g., a cryovial versus broth or agar media), and reduce the chances of mutation events. Of course, mutations and loss of mobile genetic elements can still occur, and this phenomenon was observed in Staphylococcus aureus strains that lost the mecA gene during longterm preservation at −80°C (15). Similar to those with other preservation methods, survival rates after freeze-drying vary with species. Evaluating microorganisms over a 10-year period, Miyamoto-Shinohara et al. found that survival rates after freeze-drying for Brevibacterium spp. and Corynebacterium spp. approached 80%, whereas those for Streptococcus mutans decreased to 20% after 10 years (13).

Ultralow-Temperature Freezing Microorganisms can be maintained at temperatures of −70°C or lower for prolonged periods. Systems for achieving these temperatures include ultralow-temperature electric freezers and liquid nitrogen storage units. With either system, unwanted heating can occur due to the loss of electrical power or liquid nitrogen. Close observation of the system and an adequate alarm mechanism are essential, since any increase in temperature will reduce viability. In the event that the temperature does rise, restoring power and returning to the target storage temperature as quickly as possible are essential. The presence of a cryopreservative such as glycerol may reduce the risk to microorganisms upon short exposure to higher temperatures (16). If thawing does occur, there are no guidelines for rapid restoration of the storage condition. Refreezing of the sealed vials as described below may be considered. For long-term storage, temperatures below −130°C are recommended for fastidious cells, such as fungal hyphae and protozoa. Cellular activity and chemical reactions cease at these low temperatures, but at −70°C they may still continue to a limited extent. Hence, for long-term cryopreservation of certain organisms, storage in liquid nitrogen (−196°C) or liquid nitrogen vapor (−150°C) is recommended (3).

11. Procedures for the Storage of Microorganisms n

163

Storage Vials

Cryoprotective Agents

Storage vials must be able to withstand very low temperatures and maintain a seal for their contents. Plastic (polypropylene) or glass (borosilicate) tubes may be used. Plastic vials with screw tops and silicone washers are much easier to use than glass vials that must be sealed with a flame and then scored and broken open. Several commercial suppliers stock acceptable vials, e.g., Fisher Scientific Products (Pittsburgh, PA), VWR Scientific (Radnor, PA), Wheaton Science Products (Millville, NJ), and Becton Dickinson and Co. (Franklin Lakes, NJ). Vials come in a variety of sizes. Half-dram vials are available from several suppliers and can be conveniently packaged in a 12-by-12 grid so that 144 vials are stored in one box or layer.

To protect microorganisms from damage during the freezing process, during storage, and during thawing, cryoprotective agents are often added to the culture suspension. Whereas most bacteria, fungi, and viruses survive better with such additives, studies have shown that cryoprotective agents significantly damage others. The reader is referred to detailed references for specifics (Table 1) (3, 6). Rapid freezing without additives may still be acceptable for the long-term survival of protozoa, although freeze-drying may be preferred. There are two types of cryoprotective agents: those that enter the cell and protect the intracellular environment and others that protect the external milieu of the organism.

TABLE 1

Common procedures for preservation of microorganisms

Organism group Streptococci

Mycobacteria

Spore-forming bacteria

Other Grampositive bacteria

Gram-negative bacteria

Storage method

Cryopreservative

Freezing Ultralow-temp freezing Lyophilization Freezing Ultralow-temp freezing Lyophilization Transfer Immersion in mineral oil Drying Freezing Ultralow-temp freezing Lyophilization Transfer Immersion in mineral oil Freezing Ultralow-temp freezing

Skim milk Skim milk Skim milk Skim milk Skim milk Skim milk None None None Glucose Skim milk, glycerol Skim milk, lactose None None Sucrose, glycerol Skim milk, sucrose, glycerol Skim milk, sucrose None None Sucrose, lactose Sucrose, lactose, glycerol Skim milk, sucrose, lactose None None None Soil, silica gel Glycerol, DMSO Glycerol, sucrose, DMSO, skim milk None Nutrient medium Glycerol, DMSO, skim milk Nutrient medium Blood, nutrient broth with DMSO or sucrose Blood, nutrient medium with DMSO or glycerol Nutrient medium SPGA SPGA

Lyophilization Transfer Immersion in mineral oil Freezing Ultralow-temp freezing Lyophilization

Filamentous fungi

Yeasts

Protozoa

Transfer Immersion in mineral oil Storage in distilled water Drying Ultralow-temp freezing Lyophilization (sporeformers) Storage in distilled water Drying Ultralow-temp freezing Lyophilization Freezing

Ultralow-temp freezing

Viruses

Transfer Ultralow-temp freezing Lyophilization

Storage temp (°C)

Storage duration (yr)

−20 −70 to −196 4 −20 −70 to −196 4 Room temp 4 Room temp −20 −70 to −196 4 Room temp 4 −20 −70 to −196

0.2 0.2–1 0.5–30 3–5 3–5 16–30 0.2–1 1 1–2 1–2 2–30 30 0.2–0.3 0.6–2 1–3 1–30

4 Room temp 4 −20 −70 to −196

30 0.1–0.3 1–2 1–2 2–30

4

30

4 to 25 Room temp Room temp Room temp −70 to −196 4

2–10 1–40 1–10 1–4 2–30 2–30

Room temp Room temp −70 to −196

1–2 1–2 2–30

4 −20 to −40

2+

−70 to −196

4 −70 to −196 4

0.5 1–30 6–10

164 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

Glycerol and dimethyl sulfoxide (DMSO) are most often used for the former; sucrose, lactose, glucose, mannitol, sorbitol, dextran, polyvinylpyrrolidone, polyglycol, and skim milk are used for the latter. Combinations of agents as well as detergents (e.g., Tween 80 and Triton WR 1339), other carbohydrates (e.g., honey), and calcium lactobionate have also been used. The most universal cryoprotectant is DMSO; however, the optimal cryoprotectant often varies with the microorganism. For example, glycerol appears to be best suited for the preservation of bacteria. A current and comprehensive review of protectant additives used in the cryopreservation of microorganisms is provided by Hubalek (17). Glycerol is typically added at a concentration of 10% (vol/vol), and DMSO is added at 5% (vol/vol). Prior to use, glycerol is sterilized by autoclaving. Once prepared, it can be stocked at room temperature for months. DMSO must be filter sterilized and can be stored in open containers for only 1 month prior to use. Of the external products, skim milk is the most often used. Dehydrated skim milk is purchased from medical product suppliers, e.g., Becton Dickinson and Co. and Hardy Diagnostics (Santa Maria, CA). It is autoclaved and used in a final concentration of 20% (wt/vol) in distilled water (3). This is double the concentration suggested by the manufacturers if the intent is to make a reconstituted equivalent of regular milk.

Preparation of Microorganisms for Freezing Microorganisms are inoculated into a medium that adequately supports maximal growth. Cultures are allowed to mature to the late growth or stationary phase before being harvested. Broth specimens are centrifuged to create a pellet of microorganisms. The pellet is withdrawn and resuspended in 2 to 5 ml of broth with the appropriate concentration of cryoprotectant additive. For agar specimens, broth containing the cryoprotectant is placed on the surface of the agar. The surface is scraped with a pipette or sterile loop to suspend microorganisms, and then the broth mixture is pipetted directly into freezer vials. Alternatively, the agar surface can be scraped with a sterile loop. The microorganisms can then be transferred directly into the vial of cryoprotectant and emulsified into a final dense suspension. The volume of the aliquots to be frozen is typically 0.2 to 0.5 ml.

will coat the beads, and then the excess culture suspension is aspirated from the vial. Individual beads can then be removed from storage for reconstitution without thawing the entire sample (18).

Thawing Damage to microorganisms occurs as they are warmed from the frozen state. Critical temperatures appear to be between −40 and −5°C. Studies suggest that rapid warming through these temperatures improves recovery rates. For optimal results, stored culture vials should be warmed rapidly in a 35°C water bath until all ice has disappeared (3, 6). Once a vial is thawed, it should be opened and the organism should be transferred to an appropriate growth medium immediately. Great care must be exercised during the thawing phase, since rapid temperature changes and resulting air pressure changes inside vials can cause the vials to explode. Protective clothing and eyewear must be worn during this process. For most practical purposes in the clinical laboratory, however, thorough thawing of stored bacteria or yeast in a water bath is not practical or necessary. The frozen vial can be thawed at room temperature and plated with good results for most routine organisms. If the organism vial must be saved for reuse at a later date, one may scrape off a small portion of the frozen contents with a sterile loop or pipette tip and then inoculate the appropriate media. The vial may be returned to frozen storage immediately without thawing and may be reused at a later date with limited damage to the organism.

Specialized Storage Systems Computerized technology can facilitate the organization and database management of large storage collections. Some laboratories have developed their own tracking system, while other products are commercially available, e.g., Freezerworks (Dataworks Development, Inc., Mountlake Terrace, WA) and FreezerPro (Ruro Inc., Frederick, MD). These kinds of software allow the user to enter data regarding each sample in a repository, and the data are stored in a highly organized, searchable fashion. Radio-frequency identification (RFID) labeling may also be used for compact storage of information associated with each sample, e.g., ColdTrack vials (BioTillion LLC, Skillman, NJ) or FreezerPro RFID kit.

Freezing Method The American Type Culture Collection (ATCC) recommends slow, controlled-rate freezing at a rate of 1°C per min until the vials cool to a temperature of at least −30°C, followed by more rapid cooling until the final storage temperature is achieved (3). This protocol is best achieved using a controlled-rate freezer. As an alternative, the ATCC suggests the use of an insulated freezing chamber, such as one commercially available or simply an appropriate polystyrene box packed with insulating materials. Studies in the 1970s showed that uncontrolled-rate freezing may be acceptable for most organisms and is much less expensive or laborintensive (6). When organisms are stored in liquid nitrogen, however, it is still recommended that vials be placed initially in a −60°C freezer for 1 h and then transferred into the liquid nitrogen. When organisms are stored permanently at −60 to −70°C, the vials can be placed directly into the freezer. Vials with small glass beads or plastic beads can also be used for freezing microorganisms, e.g., CryoBank (Copan Diagnostics, Murrieta, CA) or Microbank (Pro-Lab Diagnostics, Round Rock, TX) systems. The culture suspension

Freeze-Drying (Lyophilization) Freeze-drying is considered to be the most effective way to provide long-term storage of most bacteria, yeasts, sporulating molds, and viruses. Better preservation occurs with freeze-drying than with other methods because freezedrying reduces the risk of intracellular ice crystallization, which compromises viability. Removal of water from the specimen effectively prevents this damage. Among bacteria, the relative viability with lyophilization is greatest with Gram-positive bacteria (sporeformers in particular) and decreases with Gram-negative bacteria (6, 19), but overall, the viability of bacteria can be maintained for as long as 30 years. In addition, large numbers of vials of dried microorganisms can be stored with limited space, and organisms can be easily transported long distances at room temperature. The process combines freezing and dehydration. Organisms are initially frozen and then dried by lowering the atmospheric pressure with a vacuum apparatus. Freezedrying has been extensively reviewed in the past (5), and

11. Procedures for the Storage of Microorganisms n

the required equipment includes a vacuum pump connected in line to a condenser and to the specimens. Specimens can be connected individually to the condenser (manifold method) or can be placed in a chamber where they are dehydrated in one larger air space (chamber or batch method). Alexander et al. and Heckly have both published detailed descriptions of equipment options (3, 5).

Storage Vials Glass vials are used for all freeze-dried specimens. When freeze-drying is performed in a chamber, double glass vials are used. In the chamber method, an outer soft-glass vial is added for protection and preservation of the dehydrated specimen. Silica gel granules are placed in the bottom of the outer vial before the inner vial is inserted and cushioned with cotton. For the manifold method, a single glass vial is used. For both methods, the vial containing the actual specimen is lightly plugged with absorbent cotton. The storage vial in the manifold method or the outer vial in the chamber method must be sealed to maintain the vacuum and the dry atmospheric condition. All vials are sterilized prior to use by heating in a hot-air oven.

165

Cross contamination rates vary from 0.8 to 3.3% when two different microorganisms are placed on opposite sides of the same container and are as high as 8.3 to 13.3% when microorganisms are intermingled (20). In the manifold method, a rack of individual vials is used rather than a single container. The rack is placed in a dry ice bath. After the freezing process, the vials are connected by individual rubber tubes in sequence to the condenser container filled with the dry ice-solvent mixture and to the vacuum pump. As in the method described above, the vacuum is maintained at 30 μm Hg for 18 h and then the individual vials are sealed.

Storage Individual vials need to be appropriately labeled and sorted. Storage at room temperature does not maintain viability and is not recommended. Storage at 4°C in an ordinary refrigerator is acceptable, but survival rates may be improved at temperatures of −30 to −60°C (3, 5).

Reconstitution

Research concerning cryoprotective agents has been extensively reviewed (5). In general, the two most commonly used agents are skim milk and sucrose. Skim milk is used most often for chamber lyophilization, and sucrose is used most often for manifold lyophilization. Skim milk is prepared by making a 20% (vol/vol) solution of skim milk in distilled water. The solution is divided into 5-ml aliquots and autoclaved at 116°C, with care taken to prevent overheating and caramelization of the solution. The preparation is then used in smaller volumes as described above for freezing. Sucrose is prepared in an initial mixture of 24% (vol/vol) sucrose in water and added in equal volumes to the microorganism suspension in growth medium to make a final concentration of 12% (vol/vol).

Care must be taken when opening vials for reconstitution because of the vacuum inside the vial. Safety glasses should always be worn, and vials should be covered with gauze to prevent injury if the vial explodes when air rushes in. Reconstitution should also be conducted in a closed hood to avoid dispersal of microorganisms. The surface of the vial should be wiped with 70% alcohol, and then the top of the glass vial can be scored and broken off or punctured with a hot needle. A small amount (0.1 to 0.4 ml) of growth medium is injected into the vial with a needle and syringe or a Pasteur pipette, the contents are stirred until the specimen is dissolved, and then the entire contents are transferred with the same syringe or a pipette to appropriate broth or agar media. A purity check must be done on each specimen because of the possibility of either cross contamination or mutation during the preservation process.

Preparation of Microorganisms for Lyophilization

Newer Technologies

As with simple freezing, maximum recovery of organisms is achieved by using microorganisms in the late growth or stationary phase from the growth of an inoculum in an appropriate growth medium. High concentrations of microorganisms are considered to be important. The ATCC recommends a concentration of at least 108 CFU/ml (3), and Heckly suggests a concentration of 1010 CFU/ml or higher (6).

The long-term preservation methods previously described are specifically designed for recovery of microorganisms for further cultivation. Culture-independent tests based on antigen or nucleic acid technologies are in widespread use and do not require viable microorganisms. In this regard, storage of microorganisms to preserve their antigens or nucleic acids is also important for clinical laboratories. The use of Whatman Flinders Technology Associates (FTA) matrix cards (Whatman International Ltd., Maidstone, United Kingdom) or other filter paper-based products is a novel approach for long-term storage of microbial DNA that is safe (microorganisms are inactivated), inexpensive, and fast (21, 22). Bacterial and/or fungal cell suspensions are applied directly to dry FTA paper. The FTA cards are impregnated with buffers, free radical trap and protein denaturants that lyse cell membranes on contact, entrap DNA, and protect DNA from degradation. This technology has been successfully applied to a variety of bacteria and fungi and serves as a reusable DNA archiving system. Although beyond the scope of this chapter, direct specimens such as blood can be preserved using a dry blood spot on filter paper or with a non-paper-based matrix for future antibody or nucleic acid testing to detect HIV (23–25), hepatitis B virus (23), hepatitis C virus (23, 25), Rickettsia typhi, and Orientia tsutsugamushi (26).

Cryoprotective Agents

Freeze-Drying Methods In the chamber method, inner vials with the microorganism suspension are placed in a single layer inside a stainless steel container. This container is placed in a low-temperature freezer at −60°C for 1 h. The container is then transferred to a chamber containing a low temperature bath, e.g., dry ice in ethyl Cellosolve or other solvent, and covered with a sealable vacuum top, which is connected in sequence to a condenser reservoir also filled with the dry ice-solvent mixture and to a vacuum pump. The vacuum is maintained at a minimum of 30 μm Hg for 18 h. At the same time, the outer vials are prepared by being heated in an oven overnight, filled with silica gel granules and cotton, and placed in a dry cabinet (with 95% is highly effective against viruses; isopropanol has limited effectiveness against small or nonlipid viruses. c Not available in the United States. d ?, conflicting data. b

agents can cause side effects such as anaphylactic shock in patients who have had contact with chlorhexidine, mainly patients originating from the Far East (13, 14).

Common Antiseptic Compounds Alcoholic Compounds Alcohol is the most important skin antiseptic. For centuries, alcohols have been appreciated for their antimicrobial properties. Alcohol is defined by the FDA as having one of the following active ingredients: ethyl alcohol, 60% to 95% by volume in an aqueous solution, or isopropyl alcohol, 50% to 91.3% by volume in an aqueous solution. Ethyl alcohol (ethanol) and isopropyl alcohol (isopropanol) are the alcoholic solutions most often used as surface disinfectants and antiseptic agents in health care institutions because they possess many qualities that make them suitable both for disinfection of equipment and for antisepsis of skin. They are fast acting, minimally toxic to the skin, nonstaining, and nonallergenic. Alcohols evaporate readily, which is advantageous for most disinfection and antisepsis procedures. The uptake of alcohol by intact skin and the lungs when alcohol is used topically is below toxic levels for humans (15). Some studies indicated dermal and pulmonary absorption by inhalation of vapors of isopropyl alcohol from a commercial hand rub may occur and could interfere with religious beliefs of HCWs (15, 16). However, the World Health Organization (WHO) resolved this issue in their most recent guidelines published in 2009, and Muslims, for example, are allowed to use alcoholic compounds for hand hygiene. Alcohols have better wetting properties than water due to their lower surface tensions, which along with their cleansing and degreasing actions make alcohols effective skin antiseptics. Alcoholic formulations used to prepare the skin before invasive procedures should be filtered to ensure that they are free of spores, or 0.5% hydrogen peroxide should be added (17). Although the risk of infection is minimal, the low additional cost for a spore-free product is justified. Alcohols have some disadvantages. If alcoholic antiseptics are used repeatedly, they may dry and irritate the skin.

Therefore, preparations for hand disinfection should contain emollients. The exact mechanism by which alcohols destroy microorganisms is not fully understood. The most plausible explanation for the antimicrobial action is that alcohols coagulate (denature) proteins (e.g., enzymatic proteins), impairing specific cellular functions (18). Ethyl and isopropyl alcohols at appropriate concentrations have broad spectra of antimicrobial activity that include vegetative bacteria, fungi, and viruses. In fact, their antimicrobial efficacies are enhanced in the presence of water, with optimal alcohol concentrations being 60 to 90% by volume. Alcohols (i.e., 70 to 80% ethyl alcohol) rapidly (i.e., within 10 to 90 s) kill vegetative bacteria, such as Staphylococcus aureus, Streptococcus pyogenes, Enterobacteriaceae, and Pseudomonas aeruginosa in suspension tests (17). Isopropyl alcohol is slightly more bactericidal than ethyl alcohol (18) and is highly effective against vancomycin-resistant enterococci (19). It also has excellent activity against fungi, such as Candida spp., Cryptococcus neoformans, Blastomyces dermatitidis, Coccidioides immitis, Histoplasma capsulatum, Aspergillus niger, and dermatophytes, and mycobacteria, including Mycobacterium tuberculosis. However, alcohols generally do not destroy bacterial spores. In fact, spores of Clostridium difficile are stored in ethanol in the microbiology laboratory. In addition, fatal infections due to Clostridium spp. occurred when alcohol was used to sterilize surgical instruments. Both ethyl and isopropyl alcohols inactivate most viruses with a lipid envelope (e.g., influenza virus, herpes simplex virus, and adenovirus). However, several investigators found that isopropyl alcohol had lower virucidal activity against naked, nonenveloped viruses (20). In the experiments by Klein and DeForest, 2-propanol, even at 95%, could not inactivate the nonenveloped poliovirus type 1 and coxsackievirus type B in 10 min (21). In contrast, 70% ethanol inactivated these enteroviruses (21). Neither 70% ethanol nor 45% 2propanol killed hepatitis A virus (HAV) when their activities were assessed on stainless steel disks contaminated with fecally suspended virus. Among 20 disinfectants tested, only 3 reduced the titer of HAV by greater than 99.9% in 1 min

13. Decontamination, Disinfection, and Sterilization n

(2% glutaraldehyde, sodium hypochlorite with >5,000 ppm free chlorine, and a quaternary ammonium formulation containing 23% HCl) (22). Bond et al. (23) and Kobayashi et al. (24) demonstrated that 2-propanol (70% for 10 min) or ethanol (80% for 2 min) made human plasma contaminated with hepatitis B virus (HBV) at a high titer noninfectious for susceptible chimpanzees (24). Both 15% ethyl alcohol and 35% isopropyl alcohol (25) readily inactivate human immunodeficiency virus (HIV), and 70% ethanol rapidly inactivates high titers of HIV in suspension, independent of the protein load. However, the rate of inactivation decreased when virus was dried onto a glass surface and high levels of protein were present (26). In a suspension test, 40% propanol reduced the rotavirus titer by at least 4 log in 1 min (27) and both 70% propanol and 70% ethanol reduced the release of rotavirus from contaminated fingertips by 2.7 log units. In comparison, the mean reductions obtained with liquid soap and an aqueous solution of chlorhexidine gluconate were 0.9 and 0.7 log units, respectively (28).

Chlorhexidine Chlorhexidine gluconate, a cationic bisbiguanide, has been recognized as an effective and safe antiseptic for nearly 40 years (29, 30). Chlorhexidine formulations are extensively used for surgical and hygienic hand disinfection (see previous discussion). Other applications include preoperative showers (or whole-body disinfection), antisepsis in obstetrics and gynecology, management of burns, wound antisepsis, and prevention and treatment of oral disease (plaque control, pre- and postoperative mouthwash, and oral hygiene) (29, 30). When chlorhexidine is used orally, its bitter taste must be masked and it can stain the teeth. Intravenous catheters coated with chlorhexidine and silver sulfadiazine are used to prevent catheter-associated bloodstream infections (31). In fact, an infection control program to prevent catheter-associated bloodstream infections included hand hygiene, chlorhexidine site care, and full-barrier precautions in a large clinical study in intensive care units: these interventions led to an infection rate close to zero (32). Today, chlorhexidine compounds are considered the gold standard for catheter site care (33). Chlorhexidine is most commonly formulated as a 4% aqueous solution in a detergent base. However, alcoholic preparations have been demonstrated in numerous studies to have better antimicrobial activity than detergent-based formulations (34). Bactericidal concentrations destroy the bacterial cell membrane, causing cellular constituents to leak out of the cell and cell contents to coagulate (29). The bactericidal activity of chlorhexidine gluconate against vegetative Gram-positive and Gram-negative bacteria develops within minutes. In addition, it provides a persistent antimicrobial action that prevents the regrowth of microorganisms for up to 6 h. This effect is desirable when a sustained reduction in the microbiota reduces infection risk (e.g., during surgical procedures). Chlorhexidine has little activity against bacterial and fungal spores except at high temperatures. Mycobacteria are inhibited but are not killed by aqueous solutions. Yeasts and dermatophytes are usually susceptible, although the fungicidal action varies with the species (35). Chlorhexidine is effective against lipophilic viruses (e.g., HIV, influenza virus, and herpes simplex virus types 1 and 2), but viruses such as poliovirus, coxsackievirus, and rotavirus are not inactivated (29). Unlike what occurs with povidone iodine, blood and other organic materials do not affect the antimicrobial activity of chlorhexidine significantly (36). However, inorganic anions and organic anions such as soaps are incompatible with chlorhexidine, and its activity is reduced at extreme

185

acidic or alkaline pH and in the presence of anion- and non-ion-based moisturizers and detergents. Microorganisms can contaminate chlorhexidine solutions (37), and resistant isolates have been identified. For example, Stickler found chlorhexidine-resistant Proteus mirabilis after chlorhexidine was used extensively over a long period to prepare patients for bladder catheterization (38). The chlorhexidine resistance among vegetative bacteria was thought to be limited to certain Gram-negative bacilli (such as Pseudomonas aeruginosa, Burkholderia cepacia, Proteus mirabilis, and Serratia marcescens) (39). However, genes conferring resistance to various organic cations, including chlorhexidine, have been identified in S. aureus clinical isolates (40, 41). Chlorhexidine has several other limitations. When absorbed onto cotton and other fabrics, it usually resists removal by washing. If a hypochlorite (bleach) is used during the washing procedure, a brown stain may develop (29). Long-term experience with the use of chlorhexidine has demonstrated that the incidence of hypersensitivity and skin irritation is low. However, severe allergic reactions including anaphylaxis have been reported (13, 42). Although cytotoxicity has been observed in exposed fibroblasts, no deleterious effects on wound healing have been demonstrated in vivo. There is no evidence that chlorhexidine gluconate is toxic if it is absorbed through the skin, but ototoxicity can occur when chlorhexidine is instilled into the middle ear during operative procedures. High concentrations of chlorhexidine and preparations containing other compounds (e.g., alcohols and surfactants) may damage eyes (43). In the report “Strategies to Prevent Central Line-Associated Bloodstream Infections in Acute Care Hospitals,” it has been acknowledged that no recommendations with regards to chlorhexidine gluconate antisepsis can be made for infants less than 2 months of age due to incomplete safety data in this population (44). In fact, 51% of participants in a survey who used chlorhexidine in their neonatal intensive care unit reported adverse reactions. All were skin reactions and included erythema (32%), erosions (7%), or burns (61%). Of the reported skin burns, 13 of 17 (76%) were reported to have occurred in neonates with a birth weight of less than 1,500 g (45).

Iodophors Iodophors essentially have replaced aqueous iodine and tincture as antiseptics. Iodophors are chemical complexes of iodine bound to a carrier such as polyvinylpyrrolidone (PVP) iodine (also known as povidone iodine) or ethoxylated nonionic detergents (poloxamers). These complexes gradually release small amounts of free microbicidal iodine. The most commonly used iodophor is PVP iodine. Its preparations generally contain 1 to 10% PVP iodine, which is equivalent to 0.1 to 1.0% available iodine. The active component appears to be free molecular iodine (I2). A paradoxical effect of dilution on the activity of PVP iodine has been observed. As the dilution increases, bactericidal activity increases up to a maximum and then falls (46). Commercial PVP iodine solutions at dilutions of 1:2 to 1:100 kill S. aureus and Mycobacterium chelonae more rapidly than do stock solutions (47). S. aureus can survive a 2-min exposure to full-strength PVP iodine solution but cannot survive a 15-s exposure to a 1:100 dilution of the iodophor (47). Thus, iodophors must be used at the dilution stated by the manufacturer. The exact mechanism by which iodine destroys microorganisms is not known. Iodine may react with microorganisms’ amino acids and fatty acids, destroying cell structures and enzymes (46). Depending on the concentration of free iodine and other factors, iodophors exhibit

186 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

a broad range of microbicidal activity. Commercial preparations are bactericidal, mycobactericidal, fungicidal, and virucidal but not sporicidal at the dilutions recommended for use. Prolonged contact times are required to inactivate certain fungi and bacterial spores (20). Despite their bactericidal activity, PVP iodine and poloxamer-iodine solutions can become contaminated with B. cepacia or P. aeruginosa, and contaminated solutions have caused outbreaks of pseudobacteremia and peritonitis (48, 49). In fact, B. cepacia was found to survive for up to 68 weeks in a PVP iodine antiseptic solution (50). The most likely explanation for the prolonged survival of these microorganisms in iodophor solutions is that organic or inorganic material and biofilm may mechanically protect the microorganisms. Iodophors are widely used for antisepsis of skin, mucous membranes, and wounds. A 2.5% ophthalmic solution of PVP iodine is more effective and less toxic than silver nitrate or erythromycin ointment when used as prophylaxis against neonatal conjunctivitis (ophthalmia neonatorum) (51). In some countries, PVP iodine alcoholic solutions are used extensively for skin antisepsis before invasive procedures (52). Iodophors containing higher concentrations of free iodine may be used to disinfect medical equipment. Solutions designed for use on the skin should not be used to disinfect hard surfaces because the concentrations of the antiseptic solutions are usually too low for this purpose (20). The risk of side effects, such as staining, tissue irritation, and resorption, is lower for iodophors than for aqueous iodine. Iodophors do not corrode metal surfaces (46). However, a body surface treated with an iodine or iodophor solution may absorb free iodine. Consequently, increased serum iodine levels (and serum iodide levels) have been found in patients, especially when large areas were treated for a long period (46). For this reason, hyperthyroidism and other disorders of thyroid functions are contraindications for the use of iodine-containing preparations. Likewise, iodophors should not be applied to pregnant and nursing women or to newborns and infants (53). Because severe local and systemic allergic reactions have been observed, iodophors and iodine should not be used in patients with allergies to these preparations (54). Iodophors have little if any residual effect. However, for a limited time, they may have residual bactericidal activity on the skin surface because free iodine diffuses into deep regions but also back to the skin surface (46). The antimicrobial efficacy of iodophors is reduced in the presence of organic material such as blood.

in antimicrobial resistance (57), and some of these mechanisms may account for the observed cross-resistance of laboratory isolates to antimicrobial agents (58). Consequently, concerns have been raised that widespread use of triclosan formulations in non-health-care settings and products may select for biocide resistance and even cross-resistance to antibiotics. However, environmental surveys have not demonstrated an association between triclosan usage and antibiotic resistance (59). Triclosan solutions produce a sustained residual effect against resident and transient microbiotas, which is minimally affected by organic matter. Numerous studies have not identified toxic, allergenic, mutagenic, or carcinogenic potential. Triclosan formulations may help control outbreaks of methicillin-resistant S. aureus (MRSA) when used for hand hygiene and as a bathing cleanser for patients (55). However, some MRSA isolates have reduced triclosan susceptibility. Triclosan formulations are less effective than 2 to 4% chlorhexidine gluconate when used for surgical scrub solutions; properly formulated triclosan solutions may be used for hygienic hand washing. PCMX (chloroxylenol) is an antimicrobial used in hand-washing products. It is available at concentrations of 0.5 to 3.75%. Its properties are similar to those of triclosan. Nonionic surfactants may neutralize PCMX.

Octenidine Octenidine dihydrochloride is a novel bispyridine compound which is an effective and safe antiseptic agent. The 0.1% commercial formulation favorably compared to other antiseptics with respect to antimicrobial activity and toxicological properties. In vitro and in vivo it rapidly killed both Gram-positive and Gram-negative bacteria as well as fungi (60, 61). Octenidine is virucidal against HIV, HBV, and herpes simplex virus. Similar to chlorhexidine, it has a marked residual effect. No toxicological problems have been found when the 0.1% formulation was applied according to the manufacturer’s recommendations. The colorless solution is a useful antiseptic for mucous membranes of the female and male genitals and the oral cavity (62), but its bad taste limits its use orally. In a recent observational study, the 0.1% formulation was highly effective and well tolerated for the care of central venous catheter insertion sites (63). The results of this study have also been supported by a randomized controlled clinical trial (64). Octenidine is not registered for use in the United States.

Hygienic Hand Washing and Hand Disinfection Triclosan and PCMX Triclosan (Irgasan DP-300 or Irgacare MP) has been used for more than 30 years in a wide array of skin care products, including hand washes, surgical scrubs, and consumer products. A review of its effectiveness and safety in health care settings has been published (55). A concentration of 1% has good activity against Gram-positive bacteria, including antibiotic-resistant strains, but less activity against Gramnegative organisms, mycobacteria, and fungi. Limited data suggest that triclosan has a relatively broad antiviral spectrum, with high-level activity against enveloped viruses, such as HIV type 1, influenza A virus, and herpes simplex virus type 1. However, the nonenveloped viruses proved more difficult to inactivate. Clinical strains of S. aureus with low-level resistance to triclosan have been identified, but the clinical significance of this remains unknown (56). Triclosan is added to various soaps, lotions, deodorants, toothpastes, mouth rinses, commonly used household fabrics, plastics, and medical devices. Moreover, the mechanisms of triclosan resistance may be similar to those involved

Hand hygiene is the single most important infection control measure (65). However, it remains difficult to motivate HCWs to perform this simple procedure faithfully (66). The Centers for Disease Control and Prevention (CDC) has published detailed guidelines on hand hygiene (65), and in 2006, the WHO launched a global effort to improve hand hygiene in health care facilities with a reference book published in 2009 (http://whqlibdoc.who.int/hq/2009/WHO_ IER_PSP_2009.07_eng.pdf). In-depth reviews have been published by several authors (67–69). Microorganisms on the hands can be classified into three groups (70): (i) the transient biota, which consists of contaminants taken up from the environment; (ii) the resident biota, which consists of permanent microorganisms on the skin (69); and (iii) the infectious biota. Resident bacteria, most of which are on the uppermost level of the stratum corneum, have low pathogenicity and infectivity, and persons with normal immune systems who do not have implants or foreign bodies rarely acquire infections with these organisms. The density of resident bacteria on the skin ranges between 102 and 103

13. Decontamination, Disinfection, and Sterilization n

CFU/cm2, and these resident bacteria limit colonization with more pathogenic microorganisms (i.e., colonization resistance). During their daily work, HCWs can contaminate their hands with pathogens. If they do not practice good hand hygiene, they can transmit these organisms to susceptible patients. Several studies indicated that pathogens such as S. aureus (71), Klebsiella pneumoniae (72), Acinetobacter spp., Enterobacter spp., or Candida spp. can be found on the hands of >20% of HCWs. Moreover, numerous epidemics have been traced to HCWs’ contaminated hands (73–77). The goal of hand hygiene outside the operating room is to eliminate the transient biota without altering the resident biota. Hand washing for 15 and 30 s kills 0.6 to 1.1 and 1.8 to 2.8 log units, respectively (78). However, HCWs are very busy and frequently wash their hands for less than 10 s, which is insufficient to kill the transient biota (68, 69). One major advantage of the alcohol-based hand rub is that performance with these products takes about 25% of the time required for hand washing (68, 69). Moreover, compliance with hand-washing procedures does not exceed 40% even under controlled study conditions (65). However, recent studies have shown that compliance with using the alcohol-based hand rubs exceeds that of hand washing (79). Furthermore, other studies have demonstrated that rubbing one’s hands with an alcohol-based hand rub kills bacteria and most viruses more effectively than hand washing with a medicated soap (80, 81). Of note, investigators have not determined whether the level of killing is associated with the efficacy of preventing nosocomial infections. Alcohol-based hand rubs have several other practical advantages for hand hygiene over washing with soap and water. Compared with sinks, dispensers for the alcoholbased products are inexpensive, and they can be installed at locations that are more convenient for HCWs. Furthermore, unlike sinks (82), the dispensers have not been associated with outbreaks. Given the numerous advantages of these products, CDC’s current hand hygiene guidelines recommend that health care facilities consider introducing alcohol-based hand rubs as the primary mode of hand hygiene (65). Most U.S. institutions promote hand hygiene using an alcoholic hand rub as standard of care, driven by the guidelines issued by WHO in a draft form in 2004 and finalized in 2009. The advantages of alcohols also apply in the laboratory setting. Alcohol-based hand rubs can be provided in locations near the benches. They offer personnel protection and prevent cross-contamination of laboratory specimens (83). Because alcohols are flammable, health care facilities or laboratories in the United States should consult with the fire marshall before installing dispensers. Many states have laws that prohibit placing multiple containers in emergency exits and halls. However, there are only few published reports of fires caused by these products (84, 85).

Surgical Hand Washing (Scrub) or Surgical Hand Disinfection (Rub-In) In contrast to hand hygiene outside the operating room, the surgical hand scrub aims to eliminate both transient biota and resident biota so that if the surgeon’s gloves are punctured or torn, bacteria from his or her hands do not contaminate the surgical site. Tiny holes are observed in ≥30% of surgeons’ gloves after operations, even when highquality gloves are used. Cruse and Foord found that the incidence of surgical site infection was three times higher if the surgeon’s gloves were punctured than if they were intact after the procedure (5.7 and 1.7%, respectively) (86). An experimental study demonstrated that the level of bacte-

187

rial leakage through pinholes ranged between 103 and 104 CFU (87). Recently, a clinical trial clearly demonstrated that the presence of holes in a surgical glove without adequate antimicrobial prophylaxis increases the risk of postoperative surgical site infections fourfold (88). Moreover, a persistent antimicrobial effect is required after washing or disinfection to limit bacterial regrowth underneath the gloves (89). Thus, antiseptic preparations intended for use as surgical hand preparation are evaluated for their ability to reduce the number of bacteria released from hands (i) immediately after scrubbing, (ii) after wearing surgical gloves for 6 h (persistent activity), and (iii) after numerous applications over 5 days (cumulative activity). Immediate and persistent activities are considered the most important. Guidelines in the United States recommend that agents used for surgical hand preparation should significantly reduce microorganisms on intact skin, contain a nonirritating antimicrobial preparation, have broad-spectrum activity, and be fast acting and persistent. Agents, such as chlorhexidine, that have a prolonged postexposure effect are preferred because of this theoretical advantage, but there are no data from controlled clinical trials proving that the incidence of surgical site infections is lower when this agent is used. The WHO has issued a guideline for surgical hand antisepsis (www.who.int) with an executive summary published (90, 91). Alcohol-based surgical rubs have several advantages over traditional surgical scrubs. Alcoholic preparations are more effective than any medicated soap for the surgical scrub, and they do not alter the skin as much as chlorhexidine washes do. Moreover, the water supply in an operating room could harbor Pseudomonas spp. that might contaminate the hands of surgical personnel after they perform their surgical scrub (92). Brushes, which are used during a surgical scrub, may do more harm than good, and they should be used only to clean the fingernails, not to clean the skin. Given the advantages of the alcohol-based preparations, the presurgical scrub has been replaced in many European countries by the alcohol-based surgical rubs (68), and the WHO guidelines recommend the surgical hand rubs. Alcoholic gels are frequently promoted, but most of them are significantly less effective than liquids and should not be used in the operating room (93). A very rapid protocol (1.5 min) for the surgical hand rub has been proposed and was rapidly accepted by surgeons at the author’s institution (94). The results of this investigational study were confirmed in a clinical trial (91, 95). However, few commercially available products have been successfully tested, and quite a few failed. Therefore, a 1.5-min surgical hand antisepsis is only acceptable if the product is cleared for such a short exposure to the hands. Of note, both routine hand hygiene and surgical hand preparations must balance removing unwanted bacteria from HCWs’ hands and maintaining the integrity of the HCWs’ skin because damaged skin is more likely than normal skin to become colonized with pathogenic organisms. Therefore, either hand hygiene products should contain emollients or health care facilities should provide moisturizing hand lotions that do not damage latex for their staff so that their skin does not become dry, cracked, and irritated.

Presurgical Skin Disinfection The aim of skin disinfection is to remove and kill the skin biota at the site of a planned surgical incision rapidly. However, currently available antiseptics do not eliminate all microorganisms at the incision site. In fact, coagulasenegative staphylococci can be frequently isolated even after three applications of agents such as iodine-alcohol to the

188 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

skin (96). The FDA defines a skin disinfectant as a “fastacting, broad-spectrum, and persistent antiseptic-containing preparation that significantly reduces the number of microorganisms on intact skin” (97). Spore-free alcohols are well suited for this purpose, but they lack persistent activity. Iodine is frequently added for this purpose (98). PVP iodine continuously releases free iodine, which results in a limited prolonged antimicrobial effect. Chlorhexidine, with its profound prolonged effect after application, seems to be favorable compared to PVP iodine; however, PVP excelled against chlorhexidine according to a clinical trial (99). The WHO guideline on “safe surgery” recommends PVP iodine or chlorhexidine as a reasonable choice for preoperative skin preparation (http://www.who.int/patientsafety/safesurgery/ en). A recent randomized controlled clinical trial indicates a preference for chlorhexidine-alcohol over iodophors (100), but the addition of alcohol may be the reason for the favorable effect rather than chlorhexidine, as shown in a similar trial favoring iodophor in alcohol (99). A recent study also showed that microorganisms causing surgical site infection (SSI) did not at all correlate with remaining bacteria after standardized three-step disinfection with povidone iodine-alcohol (101). A Cochrane review found some evidence that preoperative skin preparation with 0.5% chlorhexidine in alcohol was associated with lower rates of SSIs following clean surgery than alcohol-based povidone iodine paint. However, the authors stated that further studies are needed that evaluate the iodine-containing and chlorhexidine-containing solutions relevant to current practice as well as the type of solution used (alcohol versus aqueous) (102). Before a patient’s skin is prepared for a surgical procedure, the skin should be free of gross contamination (i.e., dirt, soil, or any other debris) (103). Although preoperative showering has not been shown to reduce the incidence of surgical site infections (104), this practice may decrease bacterial counts and ensure that the skin is clean (105). The antiseptics used to prepare the skin should be applied using sterile supplies and gloves or by a no-touch technique, moving from the incision area to the periphery (103). The person preparing the skin should use pressure because friction increases the antibacterial effect of the antiseptic. For example, alcohol applied without friction reduces bacterial counts by 1.0 to 1.2 log CFU compared with 1.9 to 3.0 log CFU when friction is used. In comparison, alcoholic sprays have little antimicrobial effect and produce potentially explosive vapors (106).

Decolonization The term “decolonization” when used in the context of MRSA carriers refers to measures primarily aimed at reducing the number of MRSA colonies present on the skin and mucous membranes or even at eradicating (i.e., eliminating) the microorganism completely. In contrast to medical treatment, decolonization of MRSA can also be indicated in the absence of infection. In most situations, complete MRSA eradication will be attempted; however, in many circumstances, the decreased risk of infection or decreased rates of transmission may be considered a desirable outcome. Antiseptics have long been used on MRSA-colonized intact skin for decolonization. However, until now, no randomized controlled clinical studies on the efficacy of skin antiseptics used as monotherapy have been published, and the efficacy of several antiseptic agents has been demonstrated in experimental laboratory studies only (107–109). Chlorhexidine is the most extensively investigated antiseptic agent. It has been used for many years all over the

world to decolonize the MRSA-colonized skin. As early as 1977, Davies et al. (110) noted that chlorhexidine produced a significant decrease of the local skin flora in comparison with soap baths and other antiseptic agents (PVP iodine and hexachlorophane) (111, 112). This observation was confirmed by several other studies (112, 113). In the 1980s, chlorhexidine added to shower or bath water preoperatively with the aim of preventing SSIs was investigated with inconclusive results (105, 111, 112–115). Subsequently, chlorhexidine was repeatedly used for decolonization of MRSA patients (116–118); however, these investigations conducted in the setting of an outbreak were essentially observational in nature and did not include control groups. In two randomized placebo-controlled trials of whole-body washing with chlorhexidine in addition to nasal mupirocin, the superiority of chlorhexidine could not be demonstrated (112, 119). Although MRSA may not be completely eradicated by chlorhexidine, Sandri et al. (120) observed in a 5-year prospective study that the incidence of nosocomial MRSA infections was diminished following the administration of chlorhexidine plus nasal mupirocin ointment for all patients in an intensive care ward. Furthermore, in a multicenter study, it was demonstrated that universal decolonization of all patients in intensive care units was more effective than targeted decolonization or screening and isolation in reducing rates of MRSA clinical isolates and bloodstream infection from any pathogen (121). Due to the limitations of chlorhexidine, other antiseptics have been used as part of decolonization bundles. Various case reports, but no controlled studies, are available for octenidine dihydrochloride. A significant reduction in numbers of bacteria on the skin was demonstrated for several body sites (122, 123), but side effects that led to discontinuation of treatment in 14% of patients were described (123). Further antiseptics that have been proposed for eradication of MRSA skin carriage include povidone iodine (PVP iodine) (124), polyhexanide, triclosan (125), hexachlorophane (126, 127), undecylenamidopropyltrimonium methosulfate 4%, a quaternary surfactant, and phenoxyethanol 2% (128). However, none of these drugs has been studied in controlled trials. Povidone iodine (PVP iodine) is not recommended for application to extensive body surface areas due to systemic absorption (129, 130). Triclosan 0.3% added to hand or bathing soap was able to stop a MRSA outbreak affecting a neonatal unit (125), but it has been shown that MRSA strains can develop resistance to triclosan (131–133). Results on efficacy of hexachlorophane are inconclusive, and they are derived from an outbreak setting alone (126). Since antiseptic agents may be applied to extensive surfaces of the body, depending on the indication, it is advisable to select antiseptic products with low absorption to limit the risk of adverse effects. Although MRSA carriage in the throat is not uncommon (116, 134, 135), no studies have been reported that focused on eradication of MRSA from this location. Antiseptic agents potentially suitable for oropharyngeal MRSA decolonization include octenidine, triclosan, and chlorhexidine (136–140). The main difficulty consists of achieving a sufficiently long contact time between the throat and the antiseptic agent. Mouth rinses and gargling do not ensure adequate contact times, and administration of lozenges does not lead to sufficient distribution of the antiseptic. Hence, sprays may be more suitable for oropharyngeal administration (140). However, eradication of MRSA from the throat remains a largely unresolved issue and has been treated by tonsillectomy in extreme cases (141). Difficulties

13. Decontamination, Disinfection, and Sterilization n

of oropharyngeal MRSA eradication may arise from the colonization pattern of S. aureus in tonsillar tissue. Brook et al. (135) demonstrated a higher recovery rate for S. aureus from tonsillar cores than from tonsillar surfaces.

CLEANING AND DECONTAMINATION In Europe, decontamination basically means cleaning an item to remove organic material, protein, and fat. In the United States, the term describes a cleaning step and any additional step required to eliminate any risk of infection to HCWs while they handle a device without protective attire. The CDC guideline defines cleaning as the removal of visible soil (e.g., organic and inorganic material) from objects and surfaces by using water with detergents or enzymatic products. Thorough cleaning is essential before highlevel disinfection and sterilization because inorganic and organic materials that remain on the surfaces of instruments interfere with the effectiveness of these processes. Decontamination removes pathogenic microorganisms from objects so they are safe to handle, use, or discard (142). The FDA defines the cleaning process as including all steps

TABLE 2

189

necessary to remove, inactivate, or contain contamination, beginning immediately after an item has been used for clinical purposes, continuing with the steps to decontaminate, clean, and package a device up to the first step of the sterilization process and ending with quality control tests. Regardless of regulations, cleaning is always the initial step of the decontamination process on both continents. In this chapter, we use the term “decontamination” to describe the removal of debris, blood, proteins, and most microorganisms. This process usually, but not necessarily, renders the device “safe to handle” by HCWs who are not wearing protective attire. Basic definitions are outlined in Table 2. The first step in reprocessing used medical devices is for HCWs to prevent debris from drying on the item. Research on prion diseases demonstrated that removal of debris is seriously impaired if the debris is allowed to dry on a medical device (143). Therefore, the reprocessing cycle should start as soon as possible: the item should be kept wet if delays in reprocessing are anticipated (144, 145). Cleaning can be done physically or chemically; it can also be done manually, by sonication, or with washers. In the United States, cleaning is

Definitions and terms

Term Sterilization

Disinfection

Decontamination

Standard A (closely monitored) validated process used to render a product free of all forms of viable microorganisms, including all bacterial endosporesa Elimination of most if not all pathogenic microorganisms, excluding spores

Reduction of pathogenic microorganisms to a level where items are “safe to handle” without protective attire

Technical-microbiological log CFU reduction

Comment

≥6 log CFU reduction of the most resistant spores for the sterilization process studied, achieved at the half-time of the regular cycle (ISO 14937) There is not a clear-cut defined reduction level; a minimum estimate is ≥3 log CFU reduction of microorganisms, excluding spores; common reduction is 4 to 5 log for devices; these are estimates, because there is no international standardization Elimination of debris and proteins by cleaning and/or disinfection/ sterilization process; in Europe, it is restricted to cleaning only, which achieves a minimum of ≥1 log CFU; most cleaning processes achieve 3 to 5 log CFU reduction; these are estimates, because there is no international standardization

Prions require an adapted definition because of their high resistance to any form of sterilization

Preoperative skin preparation with an alcohol-based iodine compound Hand washing (scrub): reduction of ≥1 log CFU Hand disinfection (rub-in): reduction of ≥2.5 log CFU

Antiseptic agents are handled as drugs by the FDA

Some high-level disinfectants achieve microbial reduction, including spores, similar to sterilization, if long incubation times and/or temperatures of >25°C are applied; this is called liquid sterilization by sterilants

Manual and/or mechanical cleaning (with water and detergents or enzymes), a prerequisite before disinfection or sterilization; in Europe, this term is used for cleaning the items; in the United States, it defines an item to be “safe to handle”; it may include a cleaning process but also a disinfection or even a sterilization process; the U.S. term “decontamination” refers to the HCWs’ safety; in Europe, the term is used for the item only

Antisepsis Patient related

Disinfection of living tissue or skin

HCW related

Reduction or removal of transient microbiota

a Examples of standards for sterilization methods: ethylene oxide, ISO 11135 (industrial facility use) and ANSI/AAMI ST 41 (health care facility use); moist heat, ISO 11134 (industrial facility use) and ANSI/AAMI ST 46 (health care facility use).

190 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

frequently performed manually with water and a detergent. In Europe, many countries rely primarily on washer-disinfectors that rinse items with cold water and then with warm water plus a detergent. The cycle is completed with hot water at ≥90°C. Items such as bedpans and urinals can be cleaned and disinfected by putting the items into a machine, pushing a button, and removing them after a 2- to 5-min procedure. All sterilization techniques other than steaming have been shown to fail in 1 to 40% of sterilization cycles if residual proteins and/or salts are not removed by a proper cleaning process (146). Even steam sterilization at 134°C for 18 min, recommended by the WHO to inactivate prions, can fail to prevent cross-transmission if the device does not undergo a cleaning process beforehand (143–145). For floors, surfaces, and noncritical items, cleaning with a detergent is sufficient in most situations, and a disinfection process adds little if any additional effect (147). In addition, disinfectants may interfere and even lose activity with residual proteins and debris that escape the cleaning process (147). Routine disinfection of environmental surfaces in patient care areas is recommended in the United States to add a step of additional safety in cases of unrecognized body fluids, but it is restricted to intensive care units and emergency rooms in Europe (148).

DISINFECTION Disinfection is the second critical step in reprocessing medical devices. To be effective, disinfection must be preceded by thorough cleaning and must be done properly. Staff members must check the disinfectant’s concentration regularly if it is diluted at the place of use, even if it is diluted with an electronically monitored dilution device. Failures of the valve or other critical parts of the device can result in an insufficient final concentration, which usually cannot be detected by checking either the appearance or the odor of the disinfectant. Many manufacturers provide test strips to check for the appropriate concentration. Of note, numerous outbreaks have occurred when staff members have not followed appropriate protocols (149). For example, Klebsiella oxytoca caused an outbreak after an infection control committee allowed staff members to decrease the concentration of a glutaraldehyde-based surface disinfectant because they didn’t like the odor. The outbreak stopped after staff resumed using the disinfectant at the recommended concentration (150). An outbreak of 58 cases of Mycobacterium xenopi

TABLE 3

infection occurred when instruments used for discovertebral operations were rinsed with tap water after they were disinfected (151).

Definitions and Terms (Adapted from FDA and Environmental Protection Agency [EPA] Definitions) Since the FDA regulates the most critical part of disinfection and sterilization, FDA definitions are used throughout the chapter unless stated otherwise. The most important definitions are given in Table 3.

Principles and Antimicrobial Activities of Compounds The antimicrobial spectrum of disinfectants is tested differently than that of antimicrobial agents. Microbiology laboratories that test disinfectants must know the special methods needed to accurately assess their activity. In fact, MICs are of little help because the goal of disinfection is to kill rather than inhibit the growth of microorganisms. In contrast to sterilization, but similar to antimicrobial agents, killing curves for disinfectants are not linear and the rate of log killing decreases as the inoculum concentrations decrease (i.e., as the number of CFU per milliliter decreases). Therefore, a 3-log-unit killing is more easily achieved with disinfectants if the inoculum is large, e.g., 108 CFU, than if the inoculum is 104 CFU. Most disinfectants must be inactivated before they are incubated in media or plated because bacteria do not grow in the presence of very low concentrations of a disinfectant (inhibitory effect). However, if the compound is inactivated, bacterial growth can be demonstrated. Like antimicrobial agents, some disinfectants display a postexposure effect on bacterial growth. The postexposure effect has been quantified for a variety of disinfectants. Alcohols lack a postexposure effect, but chlorhexidine, octenidine, polyhexanide, and chloramine delay regrowth after exposure for several hours (23). Low-level disinfectants destroy lipid-enveloped viruses, such as HIV, and most vegetative bacteria (Fig. 1) (23), but many disinfectants, including alcohol, are ineffective against non-lipid-enveloped or small viruses such as poliovirus. For example, isopropyl alcohol has little activity against poliovirus but >90% ethanol is very active (152). The FDA requires that the microbicidal efficacy of liquid chemical sterilants and high-level disinfectants be assessed in three different types of tests before they can be legally marketed in the United States.

FDA and EPA definitions of important terms Term

Definition

Cleaning (or precleaning)

Removal of foreign material (e.g., organic or inorganic contaminants) from medical devices as part of a decontamination process Agent that destroys microorganisms; the prefixes of terms with the suffix “-cide” (e.g., virucide, fungicide, bactericide, sporicide, and tuberculocide) indicate which microorganisms the germicide kills Germicide that when used according to the labeling kills all microbial pathogens except large numbers of bacterial endospores Germicide that when used according to the labeling kills all microbial pathogens except bacterial endospores Germicide that when used according to the labeling kills most vegetative bacteria and lipid-enveloped or medium-size viruses; such disinfectants are regulated by the EPA Lowest effective concentration of a liquid chemical germicide that achieves the microbicidal activity claimed by the manufacturer Chemical germicide that achieves sterilization

Germicide

High-level disinfectant Intermediate-level disinfectant Low-level disinfectant

Minimum effective concentration Sterilant (chemical)

13. Decontamination, Disinfection, and Sterilization n

191

FIGURE 1 Microorganisms’ resistance to disinfectants. HSV, herpes simplex virus; CMV, cytomegalovirus; RSV, respiratory syncytial virus. doi:10.1128/9781555817381.ch13.f1

1. Potency testing incorporates the EPA test requirements for registration of germicides, such as the Association of Official Analytical Chemists’ (AOAC) sporicidal test, tuberculocidal test, and use-dilution tests for S. aureus ATCC 6538, Salmonella enterica serovar Choleraesuis ATCC 10708, and P. aeruginosa; EPA virucidal tests for viruses, including poliovirus type 2 and herpes simplex virus; and FDA-recommended tests, such as total killing or endpoint analysis and comparing survivor and predicted curves. (Note: in Europe, disinfectants should have been tested by the methods defined by European Norms [EN] such as EN 1040 [bactericidal activity] and EN 1275 [fungicidal activity]). 2. Simulated-use testing involves testing the disinfectant under artificially created worst-case scenarios to determine how long instruments need to be in contact with the disinfectant if cleaning failed and the instruments are still contaminated with substantial organic matter and microbes. The instruments are contaminated with an organic load and appropriate test microorganisms (the organism depends on the level of disinfection being claimed), and the conditions of the artificially contaminated devices represent worst-case postcleaning conditions prior to exposure to the germicide. 3. “In-use” testing involves cleaning medical devices used for clinical purposes according to a facility’s operating procedures.

As noted above, the FDA includes a tuberculocidal test in its testing procedures. This test does not account for the effect of cleaning before devices are disinfected. Devices are treated with 2% horse serum (proteinaceous load) and with 105 to 106 CFU of Mycobacterium terrae or equivalent nontuberculous mycobacteria. Under these conditions, a device would need to be immersed in a disinfectant (e.g., 2.4% alkaline glutaraldehyde) for ≥45 min at ≥25°C for complete tuberculocidal killing. However, Rutala and Weber demonstrated that proper cleaning eradicates at least 4 log units of microor-

ganisms (153), and Hanson et al. showed that cleaning bronchoscopes before disinfection removed all detectable contaminants, with up to an 8-log-unit reduction in the viral load (154). Therefore, Rutala and Weber recommended that the FDA accept a standardized cleaning protocol followed by a 20-min immersion at 20°C with an FDA-approved disinfectant as adequate to kill mycobacteria (153). An updated list of low-level and intermediate-level disinfectants registered by EPA or high-level disinfectants and sterilants approved by the FDA is provided on their websites: http://www.fda .gov/MedicalDevices/DeviceRegulationandGuidance/ ReprocessingofSingle-UseDevices/ucm133514.htm and http://www.epa.gov/oppad001/chemregindex.htm.

Overview of Commonly Used Disinfectants for Devices Glutaraldehyde Among aldehydes that exhibit biocidal activity, including glyoxal, ortho-phthalaldehyde (OPA), succinaldehyde, and benzaldehydes, glutaraldehyde and formaldehyde are the most extensively studied aldehydes. In-depth reviews may be found elsewhere (20, 155, 156). In commercially available products, glutaraldehyde is the predominant aldehyde. Because it has potent and broadspectrum microbicidal activity and is compatible with many materials (including metal, rubber, and plastic), glutaraldehyde is often regarded as the high-level disinfectant and chemical sterilant of choice in many health care facilities. Glutaraldehyde-based formulations are most commonly used for high-level disinfection of medical equipment such as endoscopes, transducers, dialysis systems, and anesthesia and respiratory therapy equipment (156). The mechanism of action is complex and is related to alkylation of sulfhydryl, hydroxyl, carboxy, and amino groups in the cell wall, cell

192 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

membrane, nucleic acids, enzymes, and other proteins of microorganisms. The biocidal activities of glutaraldehyde solutions are dependent on a variety of variables, such as pH, temperature, concentration at the time of use, the presence of inorganic ions, and the age of the solution (155). Aqueous solutions of glutaraldehyde are usually acidic and are not sporicidal in this form. Therefore, they need to be activated by adding an alkalinizing agent. These activated solutions, however, rapidly lose their activity because glutaraldehyde molecules polymerize at an alkaline pH. Therefore, the shelf life of such solutions is limited to 14 days unless the manufacturer recommends otherwise. To overcome this problem, some manufacturers have developed novel formulations with longer shelf lives (e.g., activated dialdehyde solutions containing 2.4 to 3.5% glutaraldehyde with a maximum reuse life of 28 days). The activities of disinfectants increase as the temperature rises. Among eight disinfectants tested, glutaraldehyde was found to be the chemical most strongly affected by temperature (157). Some stable acid glutaraldehydes may be used at temperatures of 35 to 55°C at concentrations below 2%. Glutaraldehyde retains its activity in the presence of organic matter. A standard 2% aqueous solution of glutaraldehyde buffered to pH 7.5 to 8.5 is bactericidal, tuberculocidal, sporicidal, fungicidal, and virucidal. It rapidly kills both Gram-negative and Gram-positive vegetative bacteria. Longer exposure times are required to inactivate spores and mycobacteria. Spores of Bacillus and most Clostridium spp. are generally destroyed by 2% glutaraldehyde in 3 h, whereas spores of Clostridium difficile are eliminated more rapidly (158). In contrast, Cryptosporidium parvum oocysts remained viable and infectious after 10 h in a 2.5% glutaraldehyde solution (159). Several investigators have questioned glutaraldehyde’s ability to inactivate mycobacteria. For example, Rubbo et al. (160) demonstrated that glutaraldehyde more slowly inactivated M. tuberculosis than did alcohols, formaldehyde, iodine, and phenol. Ascenzi (155) showed in the quantitative suspension test that 2% glutaraldehyde killed only 2 to 3 log units of M. tuberculosis in 20 min at 20°C. Similarly, Collins (161) reported that glutaraldehyde could not completely inactivate a standardized suspension of M. tuberculosis within 10 min. Nontuberculous mycobacteria such as Myobacterium avium, Myobacterium intracellulare, and Mycobacterium gordonae are more resistant to inactivation than M. tuberculosis (162). These and other data suggest that 20 min (at 20°C) is the minimum exposure time needed to reliably inactivate tuberculous and nontuberculous mycobacteria by 2% glutaraldehyde, provided that the contaminated item has been thoroughly cleaned before disinfection (20, 163). Glutaraldehyde-resistant mycobacteria have been isolated from endoscope washer-disinfectors (164, 165; see “Endoscopes” below). The virucidal activity of glutaraldehyde extends to the nonenveloped (hydrophilic) viruses, which are generally more resistant to disinfectants than are the enveloped (lipophilic) viruses. Numerous viruses were documented to be inactivated, including HIV, HAV, HBV, poliovirus type 1, coxsackievirus type B, yellow fever virus, and rotavirus (21, 155). The disadvantages of glutaraldehyde include the fact that it coagulates blood and can fix proteins and tissue to surfaces (20, 166). In addition, glutaraldehyde has a pungent and irritating odor, and its vapor at the level of 0.2 ppm irritates the eyes, throat, and nose. HCWs exposed to glutaraldehyde can develop allergic contact dermatitis, asthma, rhinitis, and epistaxis. Measures that may minimize employee exposure include covering immersion baths with tight-fitting lids, improved ventilation, ducted exhaust hoods or ductless fume hoods with vapor absorbents,

personal protective equipment, and appropriate automated machines for endoscope disinfection (20, 167). Due to dilution, glutaraldehyde concentrations commonly decline during use in manual and automatic baths used for endoscopes (166). Test strips should be used to ensure that the glutaraldehyde concentration has not fallen below 1 to 1.5%. Equipment disinfected with glutaraldehyde and rinsed inadequately has caused serious clinical complications, including proctocolitis (colonoscopes) (168, 169) and keratopathy (ophthalmic instruments). Because the infectivity of prions can be stabilized when instruments are treated with formaldehyde before they are autoclaved (170), aldehydes are no longer recommended for disinfecting endoscopes in some European countries (e.g., France) (see “BSE and vCJD” below).

OPA A 0.55% OPA solution has been approved as a high-level disinfectant by the FDA and by agencies in other countries. However, different countries or areas have set different exposure times for a 0.55% solution of OPA at 20°C to achieve high-level disinfection: 12 min in the United States, 10 min in Canada, and 5 min in Europe, Asia, and Latin America. Compared with glutaraldehyde, OPA has several advantages: (i) it does not require activation; (ii) it is compatible with many materials (i.e., similar to glutaraldehyde); (iii) it is more stable during storage and reuse as well as at a wide pH range of 3 to 9; (iv) it has low vapor properties; (v) its odor is barely perceptible; (vi) it is more rapidly mycobactericidal than glutaraldehyde in vitro and has good activity against glutaraldehyde-resistant strains at longer exposure times (171). However, 0.5% OPA is slowly sporicidal and does not inactivate all spores within 270 min of exposure (172). In addition, OPA stains proteins, skin, clothing, and instruments. OPA vapors may irritate the respiratory tract and eyes. At present, the effects of long-term exposure and safe exposure levels are not well defined. Therefore, OPA must be handled with appropriate safety precautions (i.e., gloves, fluid-resistant gowns, and eye protection), and it must be stored in containers with tight-fitting lids. If additional studies corroborate OPA’s advantages, this compound may replace glutaraldehyde for many uses, especially endoscope disinfection. The new agent appears to be particularly useful in washer-disinfectors, where glutaraldehyde-resistant mycobacteria have emerged (165, 172).

Formaldehyde Formaldehyde and its condensates are reviewed in depth elsewhere (173). Formaldehyde in aqueous solutions or as a gas has been used as a disinfectant and sterilant for many decades. Its use in the health care setting, however, has sharply decreased for several reasons. The irritating vapors and pungent odor produced by formaldehyde are apparent at very low levels (6 log units of vegetative bacteria, excluding spores (216). The manufacturer’s data sheets indicate good compatibility of the compound with humans, the environment, and various materials. A commercial product, available in Europe, can be used to disinfect instruments and endoscopes. Peroxygen compounds have proven efficacy against bacteria, bacterial spores, fungi, and a broad spectrum of viruses. A 1% concentration of a new commercial formulation containing peroxygen achieved a 105-fold killing of B. subtilis in 2 to 3 h in the absence of blood, but killing was poor in the presence of blood (217). Moreover, several investigators have found that peroxygen has poor mycobactericidal activity (164, 218). Besides other applications, these compounds may be suitable for disinfecting laboratory equipment and workbenches. Superoxidized water is prepared at the point of use by the electrolysis of NaCl solution, which generates hypochlorous acid and a mixture of radicals with strong oxidizing properties (219). Freshly generated solutions rapidly destroy bacteria, including spores and mycobacteria, fungi, and viruses in the absence of organic loading (220). A commercial adaptation of this process (i.e., Sterilox) has been marketed in Europe since 1999 and recently was approved by the FDA (see “Endoscopes” below) (219). Because Sterilox solutions are unstable, they should be used only once for high-level disinfection. Some investigators have claimed that superoxidized water is compatible with instruments and that it does not damage the environment, irritate the respiratory tract and skin, or corrode metal. However, others have reported that superoxidized water damages flexible endoscopes. Further studies are needed to explore the use of this new disinfectant in clinical settings. Metals such as copper and silver ions inactivate a wide variety of microorganisms (221). Although further work is required to explore their use in health care, they currently are used to disinfect water and to prevent infections associated with medical devices (e.g., intravascular catheters impregnated with silver sulfadiazine). For example, coppersilver ionization systems are successfully used to minimize legionella colonization in water systems (222). Surfacine is a new silver-based surface germicide that may be applied to inanimate or animate surfaces. Surfacine immediately eliminates microorganisms from surfaces and also has longterm residual activity (223, 224).

Emergence of Resistance to Biocides Microorganisms rarely become resistant to disinfectants. However, frequent use of sublethal concentrations of

196 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

disinfectants can select for resistant strains (225–227). Mechanisms of resistance include acquisition of resistance plasmids, changes in the cell membrane (e.g., chlorhexidine in Pseudomonas stutzeri), capsule formation (Klebsiella spp.), and activation of the norA efflux pump (S. aureus). A large proportion of household soaps now contain antibacterial agents (up to 45% in one study), which may increase the probability that resistant bacteria will emerge (228). Multiple outbreaks have been associated with soaps containing antibacterial agents such as chlorhexidine, hexetidine solution, or chlorxylenol (225–227). However, the concentrations of biocides used in the health care setting are much higher than the minimum biocidal concentrations in vitro. Therefore, resistance has not become a major problem in the clinical setting to date. However, a recent study demonstrated high-level resistance to glutaraldehyde (229). Readers desiring more information about disinfectants and antiseptics (6, 230) and resistance to these agents should read several excellent articles (6, 230–232).

Inactivation of Emerging Pathogens and AntibioticResistant Bacteria New and emerging pathogens such as the causative agent of vCJD, noroviruses, SARS and Middle East respiratory syndrome coronaviruses, avian and swine influenza viruses, hypervirulent C. difficile, Panton-Valentine leukocidin-producing S. aureus, Enterobactericeae producing extended-spectrum beta-lactamases and/or carbapenemases, and drug-resistant nonfermenting Gram-negative bacilli threaten the public health. Only limited data exist regarding the susceptibility of emerging pathogens to commonly used disinfectants or sterilants. Surrogate microbes have been studied for some pathogens. Examples include feline calicivirus for noroviruses, vaccinia virus for variola virus, and Bacillus atrophaeus (formerly B. subtilis) for Bacillus anthracis (233). Other infectious agents that cannot be evaluated by standard testing procedures (e.g., hepatitis C virus [HCV]) have been tested by alternative methods, such as PCR. With the exception of prions, there is no evidence that emerging pathogens are less susceptible to approved standard disinfection and sterilization procedures than are comparable classical pathogens. Standard disinfection and sterilization procedures for patient care equipment as recommended in guidelines and in this chapter are adequate to disinfect or sterilize instruments or devices contaminated with blood and other body fluids (234). Hospital disinfectants registered by the EPA, other than one peroxygen compound, do not have specific claims for activity against noroviruses. Because noroviruses are nonenveloped, most quats do not have significant activity against them. Phenolic-based preparations have been found to be active in vitro against a surrogate virus of this group. However, concentrations two- to fourfold higher than those recommended for routine use by manufacturers may be required. In the event of a norovirus outbreak, the CDC recommends using a hypochlorite solution (minimum chlorine concentration of 1,000 ppm) to decontaminate hard, nonporous, environmental surfaces (http:// www.cdc.gov/norovirus/index.html). SARS coronavirus and avian influenza virus are inactivated by sodium hypochlorite and a commercially available peroxygen compound (235); phenolic compounds and quats are less effective. A sporicidal germicide is required to efficiently eliminate C. difficile spores. In a recent study, glutaraldehyde (2%), peracetyl ions (1.6%, equivalent to 0.26% peracetic acid), and acidified nitrite demonstrated biocidal activity against C. difficile spores (236). Hypochlorite-based disinfectants have been used to disinfect environmental surfaces in areas with

ongoing transmission of C. difficile. Daily disinfection with bleach wipes with 0.55% active chlorine reduced hospitalacquired C. difficle infections on units with high endemic incidence by 85% (237). Recent outbreaks with virulent strains may require more focus on environmental cleaning and disinfection (238, 239). There is some evidence that sporulation of C. difficile can be enhanced by contact with non-chlorine-based cleaning agents in subinhibitory concentrations (240). There are no data demonstrating that disinfectants used at recommended contact conditions and concentrations are less effective against antimicrobial-resistant bacteria than against antimicrobial-susceptible bacteria (234). Inactivation of prions, including those causing vCJD, is discussed below.

Mode of Application of Chemical Disinfectants Cleaning and Disinfecting Surfaces and Floors In general, the environment is not a primary reservoir for nosocomial pathogens. However, in some cases, environmental contamination may be important. Recent examples include respiratory syncytial virus (241) and the SARS coronavirus (242). The CDC’s recent guidelines for environmental infection control in health care facilities recommend using an EPA-registered hospital detergent/disinfectant designed for general housekeeping purposes in patient care areas, especially in intensive care units, operating theaters, and emergency rooms, where blood, body fluids, or multidrug-resistant organisms may have contaminated surfaces (148). A one-step process is adequate in most areas, but a rinse step is necessary in nurseries and neonatal intensive care units, especially if a phenolic agent was used (243). Products with quats allow cleaning and disinfecting in one step, but residual quats on the surface may result in sticky, smeary surfaces. Other products may require a two-step approach (a cleaning step and a disinfection step), doubling the workload. “High-touch” surfaces (e.g., doorknobs, bed rails, and light switches) should be disinfected more frequently than “minimal-touch” surfaces. A simple detergent is adequate for cleaning surfaces for other patient care areas and in non-patient-care areas. Cleaning with a detergent is much more important than adding a disinfectant to the solution. In fact, several studies found that adding a disinfectant did not prolong the reduction in bacterial load on surfaces (146). Routine disinfection of environmental surfaces is necessary for all areas with patients in contact isolation (e.g., patients infected with MRSA). Twice-daily disinfection may be necessary to control an outbreak with vancomycin-intermediate S. aureus (239, 244). In rare situations, routine disinfection of surfaces and floors is crucial: when cases of norovirus or clusters with C. difficile or MRSA are detected, an immediate switch from cleaning floors and surfaces to using a highly active disinfectant is warranted. Several studies demonstrate a correlation between contaminated surfaces and clinical cases (245–247). When patients with suspected norovirus infection vomit, immediate disinfection of the vomitus with highly concentrated bleach or an oxygen-release compound is crucial. Norovirus is highly contagious; in fact, 100 virions are sufficient to induce infection, but >106 virions are shed by infected patients. The conventional disinfection methods may be limited by reliance on the operator to ensure appropriate selection, formulation, distribution, and contact time of the agent. “No-touch” automated room disinfection (NTD) systems have been introduced with the goal to reduce these problems. Available NTD systems include hydrogen peroxide (H2O2) vapor systems, aerosolized hydrogen peroxide, and

13. Decontamination, Disinfection, and Sterilization n

ultraviolet radiation. The more frequently described H2O2 vapor systems and the aerosolized hydrogen peroxide require isolation of air vents and doors of the room to be decontaminated. Peroxide will be either aerosolized or vaporized. There is some evidence that the use of H2O2 vapor systems reduces transmission of pathogens that are associated with the environment in the endemic as well as epidemic setting (248–252). However, the cost-effectiveness of NTD systems will be affected by the degree to which they reduce transmission and requires further evaluation. There are few studies disclosing the cost, but further information can be found in a review by Otter et al. (253).

197

tious agents: critical, semicritical, and noncritical (Table 4). The CDC cites this classification in its “Guidelines for Environmental Infection Control in Health-Care Facilities” (http://www.cdc.gov/hicpac/pdf/guidelines/eic_in_hcf_ 03.pdf), as does the FDA for approval of sterilants and highlevel disinfectants (see http://www.fda.gov/MedicalDevices/ DeviceRegulationand Guidance/ReprocessingofSingleUseDevices/ucm133514.htm). Most infection control professionals worldwide use this classification as well. However, this simple classification does not work perfectly for all devices. Even the definition of sterilization as the absence of any viable microorganisms must be revised to address the prions responsible for CJD and vCJD (258).

Disinfection of Medical Devices Classification of Devices for Reprocessing The principal goal of disinfection and sterilization of medical devices is to reduce the numbers of microorganisms on a device to a level that will prevent transmission of infectious organisms, with a considerable safety margin. The most conservative approach would be to reprocess all items and devices with overkill sterilization. Obviously, not all items must undergo the most vigorous process to eliminate any microorganisms. For example, items such as blood pressure cuffs that are used at nonsterile body sites do not need to be sterilized between patients. In contrast, only sterilization will eliminate any risk of infection for devices used in a normally sterile body site. In some cases, the best choice may be to use disposable items instead of reusable devices because reprocessing may be more expensive or does not provide the desired level of safety. The latter may apply to items in contact with neural tissue of a patient suffering from any form of CJD or with tonsils and other lymphatic tissues of persons with spongiform encephalopathy (bovine spongiform encephalopathy [BSE] or vCJD) (254–256). Therefore, devices must be classified to allow staff to define the appropriate method for disinfection and/or sterilization for each item. A classification system should balance the potential risks for transmission of infection (e.g., the infectious dose) and the resources available to achieve the necessary or desired level of antimicrobial killing. The most commonly used classification was proposed by Earle H. Spaulding in 1968 (257). He proposed three categories that are based on the devices’ potential for transmitting infec-

TABLE 4

Critical items. Items that enter normally sterile parts of the human body, such as surgical instruments, implants, or invasive monitoring devices (Table 4), are classified as “critical items.” Because items classified as critical carry the highest risk for the patient, sterilization is the preferred method for reprocessing these items. Autoclaving is the method of choice if the device is not heat labile. Alternative sterilization processes that use ethylene oxide or plasma require prolonged times, and the FDA has not approved them for use with instruments that have small dead-end lumens, which are difficult to sterilize. Liquid sterilization with a glutaraldehyde-based formulation or peracetic acid is acceptable if sterilization by one of the methods mentioned above is not feasible and the formulation and/or automated device has been cleared by the FDA. Semicritical items. Semicritical objects come into contact with mucous membranes or nonintact skin and should be free of microorganisms except spores. Intact mucous membranes generally resist bacterial spores but are susceptible to other microorganisms such as vegetative bacteria (e.g., M. tuberculosis) or viruses (e.g., HIV and cytomegalovirus). Examples of semicritical equipment include anesthesia equipment, respiratory equipment, and endoscopes. These items should be processed with a high-level disinfectant such as glutaraldehyde, stabilized hydrogen peroxide, peracetic acid, or a chlorine compound. Chlorine compounds corrode items and, therefore, are rarely used to disinfect medical devices.

Spaulding classification of devices

Clinical device Critical device

Semicritical device

Noncritical device Medical equipment

Example(s)

Medical device that is intended to enter a normally sterile environment, sterile tissue, or the vasculature Medical device that is intended to come in contact with mucous membranes or minor skin breaches Medical device that comes in contact with intact skin

Surgical instruments

High

Flexible endoscope

High, intermediate

Blood pressure cuff, electrocardiogram electrodes Examination table

Low

Intermediate or low level

Low

Low-level disinfection, sanitizer

Device or component of a device that does not typically come in direct contact with the patient

Infectious risk

Reprocessing procedure (FDA classification)

Definition

Sterilization by steam, plasma, or ethylene oxide; liquid sterilization acceptable if no other methods feasible Sterilization desirable; highlevel disinfection acceptable

198 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

Noncritical items. Noncritical items (bedside tables, crutches, stethoscopes, furniture, and floors) come into contact with intact skin only. Intact skin is a very effective barrier against microorganisms, and therefore, these items and devices do not need to be sterilized. Such items pose a low risk for direct transmission of pathogens and can usually be cleaned at the bedside or at their point of use with a low-level disinfectant. For example, HCWs can disinfect their stethoscopes by wiping the surfaces with alcohol. However, noncritical devices can contribute to the transmission of pathogens by the indirect route. For instance, up to 60% of cultures of the environment near patients colonized or infected with vancomycin-resistant enterococci are positive for this organism (259). HCWs can contaminate their hands when they touch these surfaces. If they do not practice hand hygiene, they can spread these pathogens to devices or directly to other patients. Therefore, noncritical items must be decontaminated if they are likely to be contaminated with pathogenic organisms. The FDA also developed a classification based on safety considerations and the regulations manufacturers must meet before marketing a device. Medical products are listed as class I to III products (Table 5). Simple products (e.g., a tongue depressor) are classified as medical product class I, which must meet very simple requirements before being marketed legally. Class II products (e.g., autoclaves) require a premarket notification [510(k)] demonstrating that the device is at least as safe and effective as a legally marketed device. Class III devices are those that support or sustain human life and are of substantial importance in preventing impairment of human health (e.g., a pacemaker). Due to the level of risk associated with class III devices, the FDA requires companies (section 515 of the Federal Food, Drug, and Cosmetic Act) to file a premarket approval application to obtain marketing clearance. The premarket approval application must contain sufficient valid scientific evidence documenting that the device is safe and effective for its intended use (258).

Endoscopes Reprocessing endoscopes is probably the most challenging reprocessing task in health care. Multiple reports of outbreaks associated with insufficient reprocessing techniques or defects of the endoscope have been published (Table 6). However, ample data indicate that a sufficient level of safety can be achieved even with manual disinfection of endoscopes if

the guidelines are strictly followed (260). Flexible endoscopes have intricate, sophisticated small parts that are difficult to clean but must be cleaned before they can be disinfected because organic material such as blood, feces, and respiratory secretions interfere with disinfection (261). Several studies have demonstrated the importance of cleaning in experimental studies with duck HBV, HIV, and Helicobacter pylori (262, 263). A large study in several centers in the United States found that 23.9% of the cultures of specimens from the internal channels of 71 gastrointestinal reprocessed endoscopes grew ≥10 6 CFU of bacteria and that 78% of the facilities did not sterilize all biopsy forceps (264). Other studies have documented that up to 40% of the institutions do not follow published guidelines for endoscope disinfection (167, 265, 266), and reuse of disposable endoscopic accessories is common in the United States. These items frequently are not sterilized, and reprocessing protocols are not standardized. Therefore, reused disposable items might be a source of crosstransmission (261, 267). Currently, most high-level disinfectants approved by the FDA for reprocessing endoscopes contain >2% aldehyde with or without peracetic acid (http://www.fda.gov/MedicalDevices/DeviceRegulationand Guidance/ReprocessingofSingle-UseDevices/ucm133514 .htm). However, aldehydes should only be used after completing the cleaning cycle because they may stain prions to the instruments. Endoscopes, which are semicritical items, must be immersed in ≥2% glutaraldehyde for ≥20 min to achieve the necessary level of disinfection. These parameters are sufficient to kill ≥3 log units of mycobacteria, the most-resistant vegetative bacteria. Glutaraldehyde-resistant mycobacteria have been identified (164). Several authors raised concerns that C. difficile may not be fully inactivated by standard reprocessing procedures. However, transmission of C. difficile by contaminated endoscopes has not been reported to date. Moreover, cryptosporidia withstand several hours of exposure to glutaraldehyde (159) but do not survive on dry surfaces (268). Therefore, drying before storing reprocessed items is part of the process and should not be cut to save time, e.g., in endoscopy units. The glutaraldehyde concentration in commercial cleaner-disinfectors can decrease by more than 50% after 2 weeks, which may promote the emergence of resistant bacteria (165). Higher concentrations of glutaraldehyde (3.2% instead of 2%) appear to be safe for endoscopes and achieve the required ≥3-log-unit killing with a higher margin of safety than achieved with the standard concentration (269). OPA and peracetic acid plus hydrogen peroxide can be

TABLE 5 Principles of medical device classification

Classification

Class I Class II

Class III

FDA regulation

Least regulated, requires fewest regulations Must meet federal performance standards

Implanted and life-supporting or life-sustaining devices are required to have FDA approval for safety and effectiveness

Proposed classification by Global Harmonization Task Forcea

Example(s)

None

A

Band-Aid, tongue depressor

Premarket notification [510(k)]

B

Hypodermic needles, suction equipment

C D

Orthopedic implants Pacemaker

Premarket requirements by the FDA

Premarket approval

a

Details available at http://www.imdrf.org/docs/ghtf/final/sg1/technical-docs/ghtf-sg1-n77-2012-principles-medical-devices-classification-121102.pdf.

13. Decontamination, Disinfection, and Sterilization n

199

TABLE 6 Outbreaks and pseudo-outbreaks associated with contaminated endoscopes or instruments for minimally invasive procedures No. of cases

No. of deaths

Yr of publication

K. pneumoniae

7

0

2010

P. aeruginosa

20

0

2009

P. aeruginosa

7

0

2008

P. aeruginosa

17

K. pneumoniae, Proteus vulgaris, Morganella morganii P. aeruginosa

11

0

2005

16

0

2005

P. aeruginosa

3

0

2004

P. aeruginosa

39

3

2003

P. aeruginosa

18

0

2001

M. xenopi

58

0

2001

P. aeruginosa

11

2

2000

P. aeruginosa, mycobacteria

29

0

1999

HCV

2

0

M. tuberculosis

2

M. tuberculosis (multidrug resistant) P. aeruginosa

Microorganisms

Nontuberculous mycobacteria Multiple microorganisms

Problem identified

Reference(s)

Mixed

347

Mixed

335

Infections

336

Mixed

337

Mixed

338

Mixed

261

Infections

339

Infections

5

1997

Probable defective endoscope Loose biopsy port cap in the bronchoscope Improper connection to liquid sterilization device Inappropriate disinfection of microsurgical instruments, tap water rinse after disinfection Failure of washer-disinfector, purchased without expert advice, poor maintenance Problems related to the use of Steris System 1 processor Cleaning, immersion

0

1997

5

1

23

Not reported

2006

Failure to dry the duodenoscope after resprocessing Failure of automatic endoscope reprocessor and noncompliance Failure of automatic endoscope reprocessor and noncompliance Inadequate processing and storage of a flexible bronchoscope Loose port of the bronchoscope’s biopsy channel Defective biopsy forceps

Type of outbreak

Mixed

340

Infections

151

Infections

3, 341

Mixed

271

Infections

342

Cleaning, immersion

Infections

343

1997

Cleaning, immersion

Infections

344

0

1996

Failure of washer-disinfector

345

4

0

1992

Failure of washer-disinfector

377

7

1993

Cleaning, immersion, use of tap water, poorly designed washer-disinfector

Pseudooutbreak Pseudooutbreak Infections

used to disinfect endoscopes. Because the latter might corrode some endoscopes, reprocessing staff should ensure that the manufacturer of the endoscope approves this disinfectant for reprocessing. Automated washer-disinfectors specifically for endoscopes were developed, in part, to reduce the work needed to reprocess endoscopes and to decrease the risk of human errors during manual reprocessing. These machines rinse the endoscopes, clean them in several steps, and run a full-cycle disinfection process. The time endoscopes are exposed to disinfectants is set by the machine and cannot be shortened, as it can be by busy staff manually reprocessing endoscopes. However, endoscope washers can become contaminated with pathogenic bacteria. For example, one study

346 4 (review)

found Gram-negative bacteria and/or mycobacteria in 27% of cultures of specimens obtained before the final alcohol rinse and in 10% of cultures of specimens obtained thereafter. In the same study, 37 and 27% of the manually disinfected endoscopes remained contaminated at the same time points (270). In 1992, Olympus recalled (recall no. Z-039/040-2 by the FDA) its 835 model endoscope washers because the design allowed the internal tanks and tubing to become colonized by waterborne organisms such as Pseudomonas spp. In 1999, CDC reported three outbreaks related to the Steris System 1 (271). This device is supposed to sterilize the endoscopes, but they must first be cleaned manually (272). See Table 6 for a summary of outbreaks related to endoscopes,

200 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

including those related to contaminated washer-disinfectors. Newer washer-disinfectors should continuously monitor the pressure in all channels to detect debris blocking the channels, provide adapters for all types of endoscopes, use an appropriate disinfection process with an FDA-approved disinfectant, use filtered water or sterile water for rinsing, and have a built-in automatic disinfection process. These washer-disinfectors can help staff trace problems by monitoring and documenting the disinfecting process in a manner similar to that used by autoclaves. To avoid problems, knowledgeable staff should review currently marketed machines before purchasing a washer-disinfector to ensure that the one they choose is appropriate for their needs (273). To facilitate this process, the FDA recommends that the manufacturer provide a list of all brands and models of endoscopes that are compatible with the washer-disinfector and highlight limitations associated with processing of certain brands and models of endoscopes and accessories. Preferably, the manufacturer should identify endoscopes and accessories that cannot be reliably reprocessed in the device (negative list). In addition, HCWs should be trained to use the equipment and monitored subsequently to ensure that they follow the protocol exactly. Although this is not yet mandatory, it is prudent to regularly culture the rinse water of washer-disinfectors for pathogens such as Pseudomonas spp. and Mycobacterium spp. to identify problems before clinical cases occur. In Europe, validation of the whole procedure is necessary to ensure that it complies with the requirements of the European Standard EN ISO 15883 parts 1, 4, and 5 for automated endoscope reprocessing (274). However, outbreaks may occur despite negative routine culture results (159, 268). If washer-disinfectors recycle water, residual glutaraldehyde may remain on the endoscopes. Manual reprocessing is more prone to leave residual glutaraldehyde on endoscopes than are automated washer-disinfectors (275). Thus, endoscopes that are manually disinfected should be thoroughly rinsed to remove any residual disinfectant, specifically glutaraldehyde. Patients exposed to residual glutaraldehyde can develop colitis (168, 169). Reprocessed endoscopes should be stored vertically (to facilitate drying) in a cabinet (to protect them from dust and secondary contamination). Drying cabinets with a heat fan, which keep the endoscope dry in a cleanair environment, are available. Reprocessed endoscopes that are stored for days or weeks before use probably should be reprocessed again, or alternatively, the channels should be rinsed with spore-filtered alcohol (70% g%) if this agent is compatible with the instrument. In France, reprocessing is mandatory if the reprocessed endoscope has not been used within certain time limits. However, the necessity of these precautions has not been established. Guidelines for infection prevention and control in flexible endoscopes have been updated (167) and should be consulted before choosing a method and/or disinfectant for reprocessing. A checklist adapted from the FDA recommendations may help staff reprocessing endoscopes avoid errors (http://www.fda.gov/ MedicalDevices/Safety/AlertsandNotices/PublicHealth Notifications/ucm062282.htm) (Table 7). An updated list of sterilants and high-level disinfectants approved by the FDA in a 510(k) with general claims for processing reusable medical and dental devices can be found on the FDA website (http://www.fda.gov/MedicalDevices/DeviceRegulationandGuidance/ReprocessingofSingleUseDevices/ucm133514.htm). Of note, more than 20% of all damage to endoscopes is associated with disinfecting agents. Therefore, staff members who reprocess these items must ensure that the instruments and the disinfectant are compatible (267). More detailed information is available in the

TABLE 7 Checklist to avoid reprocessing errors of endoscopesa 1

All staff must comply with the manufacturer’s instructions for cleaning endoscopes

2

Determine whether your endoscope is suitable for reprocessing in an automatic washer-disinfector, which is the preferred method

3

Compare the reprocessing instructions provided by the endoscope and washer-disinfector manufacturers and resolve any conflicting recommendations

4

Follow the instructions provided by the manufacturers of the endoscopes and the chemical germicides

5

Consider drying endoscopes with alcohol

6

Monitor adherence to the protocols for reprocessing endoscopes

7

Provide comprehensive, intensive training for all staff reprocessing endoscopes; keep records of persons attending training

8

Endoscopes sent for repairs should be labeled as “contaminated equipment for repair”

9

Implement a comprehensive quality control program

a Adapted from the FDA (http://www.fda.gov/MedicalDevices/Safety/AlertsandNotices/PublicHealthNotifications/ucm062282.htm).

new CDC guideline (www.cdc.gov/hicpac/Disinfection_ Sterilization/acknowledg.html).

Dental Equipment Critical and semicritical dental instruments should be sterilized; if they will not be used immediately, they should be packaged before they are sterilized. All high-speed dental handheld pieces should be sterilized routinely between patients. Handheld pieces that cannot be heat sterilized should be retrofitted to attain heat tolerance; if this is not feasible, they should not be used. The adequacy of sterilization cycles should be verified by periodically (e.g., at least weekly) including a biological indicator with the load. This recommendation is rarely followed in Europe (276). In fact, 33% of British dental practices do not have a policy on general disinfection and sterilization procedures and only 3% own a vacuum autoclave (277), and 52.9% of the Irish dentists who responded to a survey did not autoclave their dental handpieces and only 44.7% disinfected impressions before sending them to the laboratory (278). Environmental contamination can be a problem in dental offices (279). For example, Legionella spp. can contaminate the air-water syringes and high-speed outlets. Water used on patients infected with bloodborne pathogens can contaminate surfaces and equipment. In fact, Piazza et al. found that more than 6% of samples from workbenches, air turbine handheld pieces, holders, suction units, forceps, and dental mirrors were positive by PCR for HCV (280). Therefore, infection control issues, particularly in regard to HCV and HBV, may be more important in dentistry than has been appreciated previously. The CDC and the American Dental Association (ADA) have published guidelines for infection control in dental settings (281, 282). The ADA recommends that

13. Decontamination, Disinfection, and Sterilization n 201

metal and porcelain equipment be immersed in glutaraldehyde or exposed to this disinfectant, that removable dentures and acrylic or porcelain be disinfected with iodophors or chlorine compounds, and that wax rims or bite plates be disinfected with a spray containing iodophors. Additional information can be found on the website of the ADA (http:// catalog.ada.org/ProductCatalog/601/OSHA-InfectionControl/The-ADA-Practical-Guide-to-Effective-InfectionControl/P692).

that are immersed longer than recommended will not be damaged. The latter is important because staff might forget to remove instruments, for example, during weekends or night shifts. Prolonged exposure to a disinfectant may damage the instrument. Staff should also consider the toxicity, odor, compatibility with other compounds, and residual activity of disinfectants before choosing them (Table 8). Advice from health care professionals of different institutions is very helpful to learn about their experience and to uncover problems such as interactions with detergents, unexpected coloring, odors, and, last but not least, emotions elicited. A new disinfectant used for environmental surfaces may interact with those used in the past and temporarily release unpleasant odors. Written infection control standards for environmental surfaces help to avoid incompatibilities. It is prudent to contact colleagues already using a disinfectant before introducing it in a health care facility. Once staff have identified a product that meets a facility’s needs, only strong evidence from good studies should lead to a change to a new product (e.g., the product has improved activity or works faster).

Guidelines for Choosing a Disinfectant Rutala and Weber have published guidelines for the selection and use of disinfectants and recommendations on the preferred method for disinfection and sterilization of patient care items (20, 283). The CDC issued guidelines for environmental infection control in health care facilities, including recommendations for cleaning and disinfection (148) and updated its recommendation in 2008 (www.cdc.gov/ hicpac/Disinfection_Sterilization/acknowledg.html) (142). When choosing a disinfectant, individuals responsible for infection control should review its effectiveness against the expected spectrum of pathogens (Tables 3 and 8) to ensure that it is adequate for the intended purpose. In addition, staff must ensure that the disinfectant is compatible with the devices it is intended to disinfect and that devices

Disinfection by Heat versus Immersion in Germicides Disinfection by heat has become much more common than in the past and has replaced disinfection with germicides

TABLE 8 Overview of common disinfectantsa

+

+

+

+

−

+

−

+

+

+

+ − Endoscopes

3–25%

High/CS

+

+

+

+

+

±

+

±

−

±

+

+

−

100–1,000 ppm free chlorine

High

+

+

+

+

+

±

+

+

+

+

+

+

+

Isopropyl alcohol

60–95%

Intermediate +

+

+

±

+

−

+

±

−

±

±

+

−

Glucoprotamine

1.5–4%

Intermediate +

+

+

+

+

−

+

−

−

−

+

+

−

Phenolic compounds

0.4–5% aqueous

Intermediate +

+

+

±

+

−

+

−

+

−

+

+

−

Iodophors

30–50 ppm free iodine 0.4–1.6% aqueous

Intermediate +

+

+

+

±

−

+

±

+

+

±

+

−

+ − Contact lenses + ± Selected semicritical devices + − Smallarea surfaces − − Diagnostic instruments + + Surgical instruments + − Medical equipment

Low

+

±

−

−

−

+

−

+

+

+

+

−

±

Bacterial spores

Bacteria Quaternary ammonium compounds

Toxic

Respiratory irritant

+

Residue

+

Corrosive or deleterious effect

+

Shelf life of >1 wk

High/CS

Level of disinfection

M. tuberculosis

2–3.2%

Use dilution

Fungi

Glutaraldehyde Hydrogen peroxide Chlorine

Germicide

Lipophilic viruses

Eye irritant

Application in hospitals

Skin irritant

Environmental concerns

Important characteristics Inactivated by organic matter

Small or hydrophilic viruses

Active against:

+ − Disinfection in food preparation areas and floors

a Data from references 6, 20, 149, 226, and 329. Abbreviations: CS, chemical sterilant; +, yes; −, no; ±, variable results. Efficacy of the disinfectants is based on an exposure time of less than 30 min at room temperature. Spores require prolonged exposure times (up to 10 h) unless used with a machine at higher temperatures.

202 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

for many applications in European health care facilities (284). The advantages of these devices are obvious: (i) the processes are automated and are monitored and documented in a manner similar to that for sterilization; (ii) microorganisms have not developed resistance to these processes; (iii) the cost per load is probably less than the cost of germicides. In addition, studies by Gurevich et al. (285) indicate that pasteurization with a germicide is more effective than pasteurization without a germicide. However, washers include a cleaning process with an average reduction of 4 log units, coupled with heat disinfection (5-log-unit killing; washerdisinfectors such as the AMSCO Reliance 430 achieve an inactivation factor of >5 log units) (2, 51), resulting in a total reduction of 8 to 9 log units. This surpasses any international requirements for high-level disinfection. Thermal disinfection has several disadvantages. First, the cost to purchase and install the equipment is much higher than for systems using a germicide. Second, considerable power is needed to heat the water. Third, some non-sporeforming microorganisms such as enterococci resist temperatures of up to 71°C for 10 min. Thus, recommendations such as those in the United Kingdom (the Department of Health requires 65°C for 10 min, 71°C for 3 min, or 80°C for 1 min) may not be adequate for these organisms (286). Medical washer-disinfectors that are intended to clean, disinfect at a low or intermediate level, and dry surgical instruments, anesthesia equipment, hollowware, and other medical devices are exempt from the premarket notification procedures in subpart E of part 807 of the chapter subject to §880.9 (287). The ISO provided standards for these processes in ISO norm 15883, which defines the standards for washer-disinfectors by heat with and without the addition of disinfectants. This organization has not defined a temperature at which these devices must work but rather allows manufacturers to choose a temperature in a given range at which their devices should operate. In the United States, hot-water pasteurization is generally performed at 77°C for 30 min (288), but few scientific data support use of a particular temperature. ISO 15883 introduces the A0 concept, which is based on the fact that a defined temperature will generate a predictable lethality effect against microorganisms. Corresponding exposure temperatures and time periods that achieve high-level disinfection can be calculated assuming the presence of particularly heat-resistant microorganisms in numbers in excess of those likely to be encountered on the medical devices to be processed. ISO 15883 introduces the term A0 for moist heat disinfection (thermal disinfection). The A0 value of a moist heat disinfection process denotes the lethality effects expressed in terms of the equivalent time in seconds at a temperature of 80°C delivered by the process to the medical device with reference to microorganisms possessing a z value of 10. Given a predefined A0, equivalent killing of microorganisms is achieved if the following formula is followed: A 0 = ∆TΣ10(T−80)/z)t, where T is the temperature in degrees Celsius, and t is time in seconds. An A0 value of 600, which can be achieved at 80°C over 10 min, 90°C over 1 min, or 70°C over 100 min, is the minimum requirement for noncritical medical devices (287, 288). An A0 value of at least 3,000, which can be achieved by exposure to hot water, e.g., at 90°C (the medical device must tolerate this temperature for >5 min), must be employed for medical devices contaminated with heat-resistant viruses such as rotavirus and HBV. An A0 value of at least 3,000 is also appropriate for high-level disinfection of all semicritical devices. The test procedure based on the A0 concept has

been highly reproducible and found to be suitable to test washer-disinfectors (289, 290).

STERILIZATION Principles, Definitions, and Terms As outlined in Table 2, sterilization is not a relative term but defines the complete absence of any viable microorganisms, including spores. However, this absence cannot be proven by current microbiological techniques (291). Therefore, sterilization can be defined as a closely monitored, validated process used to render a product free of all forms of viable microorganisms, including all bacterial endospores. To test the ability of sterilization systems to meet the latter definition of sterilization, manufacturers developed a worst-case scenario that allows the process (log killing) to be quantified and estimates the probability of process failure. Large safety margins were included in this test, which is based on the assumption that items are heavily contaminated with spores, soil, and proteins. It is important to note that, while these conditions are used for testing, in clinical practice, items that are heavily soiled should not be sterilized and such a scenario would represent a critical failure of the reprocessing cycle. Any device undergoing sterilization first must undergo an appropriate cleaning process. A manufacturer must demonstrate that a sterilizer is effective against a wide range of clinically important microorganisms before being approved by the FDA. In addition, proof of efficacy must be performed with organisms (usually bacterial spores) that have been shown to be the most resistant to the new technology. A validated and reliable biological indicator must be developed, and studies must establish that sterility will be consistently achieved when critical process parameters operate within a defined range. This assures the operator that as long as there is no operational error or equipment failure, sterility is achieved. Several guidelines are essential documents for staff needing to understand reprocessing and sterilization of medical devices. ISO 14937 provides general criteria for characterizing a sterilizing agent and for the development, validation, and routine control of a sterilization process for medical devices. ISO 11134 (moist heat) and ISO 11135 (ethylene oxide) documents describe the standards for use of these methods of sterilization in the industrial setting in the United States. The American National Standard Institute/Association for the Advancement of Medical Instrumentation (ANSI/AAMI) published adaptations of these standards for health care facilities: standard 46 (moist heat) and standard 41 (ethylene oxide) (Table 2). In Europe, EN 550, EN 554, and EN 285 define the standards for steam and ethylene oxide sterilization. ISO 14161 provides guidance that staff can use when selecting and using biological indicators and when interpreting the results of these tests. ISO 17664 specifies which information medical device manufacturers must provide so that the medical device can be processed safely and continue to function properly. Readers are referred to other publications for additional information about sterilization (6, 291, 292). Hotair sterilization does not belong to the state-of-the-art technologies, but it is still used in many countries. However, the distribution of dry heat to the instruments requires long exposure times. Temperatures of >185°C resinify paraffin, destroying the lubricating function of instruments, and higher temperatures are corrosive, resulting in loss of hardness. Therefore, hot-air sterilization has largely been replaced by better, safer, and faster technologies.

13. Decontamination, Disinfection, and Sterilization n

Monitoring Any sterilization process must be monitored by mechanical, physical, chemical, and facultatively biological methods. Before routine use, the performance of the machine should be validated with the most difficult load used at the institution to ensure safety of the process. In routine use, a printout of the physical parameters (e.g., temperature and pressure) during sterilization should be kept for documentation purposes. In addition, chemical indicators placed on the tested items change color if they are exposed to adequate temperatures and exposure times. They are inexpensive and easy to use and provide a visual indication that the item has been exposed to the sterilization process. Good clinical indicators are able to identify a sterilizer failure. However, some are too sensitive, giving false-positive results (293, 294), which may cause unnecessary recalls of adequately sterilized items. Less-sensitive chemical indicators do not detect small deviations in the process. In 1963, Bowie and Dick determined that if residual air remained in a sterilizer after the vacuum phase and there was only one package in the chamber, the air would concentrate in that package (295). They developed the Bowie-Dick test to determine whether steam penetration and air removal occurred successfully. This test does not provide information about the sterilization process. Biological indicators are the best monitors of the sterilization process. If the spores in commercially available standard biological indicators do not grow during an appropriate incubation period, the results indicate that the process was able to kill ≥106 CFU. For flash sterilization, the Attest Rapid Readout biological indicator detects the presence of a spore-associated enzyme, α-D-glucosidase, and permits staff to assess the efficacy of sterilization within 60 min (296). Staff should investigate positive biologicals because they can provide the only indication that something is wrong with the sterilization process (297). An important question is whether a load can be distributed before the final results of the biological indicator are available (i.e., parametric release). The Joint Commission on Accreditation of Healthcare Organizations standard allows the use of appropriate chemical indicators without routine use of a biological indicator. A common approach is to use the sterilized items if the physical and chemical parameters of adequate sterilization were met without awaiting the culture results from the biological indicators. In Europe, routine use of biological indicators is not required if the sterilizer has undergone testing by a validation procedure used for industrial steam sterilization (EN 285, EN 550, EN 554, or EN 556). Most sterilizers in European hospitals probably do not meet these very strict requirements (298), and consequently, biological indicators are used regularly to ensure that they are working properly. These industrial standards for validation of steam sterilization will be implemented in health care organizations, but this change is controversial because of the associated expenses. The future is likely to involve parametric release with regular validation and/or commissions of the equipment. Legal aspects will probably determine the outcome of this discussion, and lawyers are likely to accept nothing but a zero risk. However, the goal of a zero risk for contamination in central sterilization services will probably contribute to excessive health care costs. Therefore, standards for sterilization should exclude a risk for contamination after the reprocessing cycle but should avoid steps that are performed only for legal reasons.

Packaging, Loading, and Storage Items that are clean and dry should be inspected and then wrapped and packaged (or put in containers) before steriliza-

203

tion. Wrappers should allow steam or gas to penetrate into the package but should protect the items from recontamination after sterilization. For steam sterilization, muslin as the only wrapper has limitations and handling of items made of muslin leads to contamination (299). Items should be labeled with information such as expiration date, type of sterilization, and identification code for traceability.

Steam Sterilization The most reliable method of sterilization is one that uses saturated steam under pressure. It is inexpensive, nontoxic, and very reliable. Steam penetrates fabrics, and its inherent safety margin is much higher than that of any other sterilization technique. Therefore, it should be used whenever possible. Obviously, other techniques must be used for heat-sensitive items. Steam irreversibly coagulates and causes denaturation of microbial enzymes and proteins. Three parameters are critical to ensure that steam sterilization is effective: the amount of time the items are exposed to steam, the temperature of the process, and moisture. Unlike time and temperature, the moisture condition in the autoclave cannot be directly determined. The D-value determines the time required to kill 106 CFU of the spores most resistant to the sterilization process under study. Devices or instruments must reach the desired temperature, which is not necessarily identical to the temperature displayed on the autoclave’s gauge. A drop of only 1.7°C (3°F) increases the time required to sterilize an item by 48%. Without moisture, a temperature of 160°C is required for dry-heat sterilization. Dry air does not provide steam for condensation, and the heat transfer to objects is slower than when moisture is present. Pressurized steam quickly transfers energy to the sterilizer load and causes more rapid denaturation and coagulation of microbial proteins. In addition, pressurizing the steam allows one to achieve dry 100% saturated steam. Thus, there is no mist that could cause the packaging and/or the items to become wet. Residual air in the autoclave interferes with the sterilization process. The amount of air within the sterilizer can be estimated by comparing the chamber pressure with the saturated steam pressure calculated from the average chamber temperature. A measured pressure greater than the calculated saturated pressure indicates the presence of residual air in the chamber. Such monitoring devices are common in the United Kingdom. Several types of autoclaves are available: gravity displacement steam sterilization, prevacuum steam sterilization, and steam flash-pressure pulsing steam sterilization autoclaves. The sterilization process is less consistent in gravity displacement steam sterilizers than in the other sterilizers (300). For example, gravity displacement autoclaves are more likely than the other systems to leave residual air in the chamber before the steam is introduced. Prevacuum sterilizers resolved part of this problem and cut the cycle time in half. However, the effectiveness of sterilization still can be compromised by small leaks (1 to 10 mm Hg/min) in the sterilizer (291). The most current technology is the steam flashpressure pulsing steam sterilization technique. Because air leaks do not decrease the effectiveness of the process, it nearly eliminates the problem of air in the chamber and it reduces the thermal lag upon heating of the load to the desired exposure temperature (300). The process of sterilization has several cycles: conditioning, exposure, and drying. Common cycles for steam sterilization in prevacuum or flash-pressure pulsing steam sterilizers are 121°C for 15 min (121°C for 30 min in a gravity displacement sterilizer) or 132°C for 4 min. EN 554 requires

204 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

steam sterilizers to provide this temperature throughout the entire chamber within a narrow margin (0 to +3°C). Flash sterilization is an emergency process used, for example, after a surgical instrument is dropped but needs to be immediately available during a procedure (301). Unwrapped devices are exposed to pressurized steam for 3 min, usually in the operating suite, sometimes without a biological indicator. The autoclaves employed are gravity displacement sterilizers that have the problems mentioned above. If HCWs are in a hurry, they may not clean the item properly, which will prevent proper sterilization. In addition, because the items are not wrapped, they can be contaminated easily when they are transported to the operating room. Even properly wrapped sterile items can become contaminated if they are transported several times (299). Moreover, some patients have been injured by items that were flash sterilized (302). Therefore, flash sterilization is controversial and several investigators have suggested that it should be used only in emergency situations when no other device is available. Flash sterilization should not replace standard sterilization protocols (303) and should not be used to save time instead of sterilizing items by the standard methods or because the health care facility does not want to purchase an extra instrument set (103). If flash sterilization of an implantable device is unavoidable, records must be kept (i.e., load identification, patient’s name/hospital identifier, and biological indicator result) to facilitate epidemiological tracking.

Ethylene Oxide Gas Temperature- and/or pressure-sensitive items have been sterilized traditionally with ethylene oxide in a standard gas. Ethylene oxide inactivates all microorganisms, including spores, probably by an alkylation process. B. subtilis bacterial spores are among the most resistant, and therefore, these are used as a biological monitor for this process. A new rapid-readout ethylene oxide biological indicator indicates an ethylene oxide sterilization process failure by producing fluorescence, which is detected in an autoreader within 4 h of incubation at 37°C, and a color change related to a change in pH of the growth media within 96 h of continued incubation (224). The process of sterilizing items with ethylene oxide begins by adding nitrogen gas to remove air or by evacuating the chamber. Items are then exposed to ethylene oxide at 55°C (130°F). Six variable but interdependent parameters—gas concentration, vacuum, pressure, temperature, relative humidity, and time of exposure—must be controlled when ethylene oxide is used. The gas concentration cannot be measured online, limiting the extent of monitoring. Therefore, the concentration should be validated as outlined in ISO 11135. Ethylene oxide sterilization has several disadvantages. It is useful only as a surface sterilizer because it does not reach blocked-off surfaces. In addition, ethylene oxide is flammable, explosive, and carcinogenic to laboratory animals, and it requires special safety precautions. Moreover, items sterilized by ethylene oxide must be aerated for approximately 12 h to remove any traces of the gas. Thus, the entire process takes >16 h, but modern sterilizers can run at shorter cycles. Furthermore, toxic residues can be trapped in the wrapper or the items. Polyvinyl chloride and polyurethane absorb ethylene oxide readily and require long periods to dissipate the oxide. The wrapper should be a barrier against recontamination after sterilization, but it also can prevent ethylene oxide from reaching the item. Therefore, only materials with documented ethylene oxide penetration and dissipation properties should be used as wrappers.

The future of ethylene oxide in sterilization is limited, mainly due to its toxicity. However, no currently available technology, including plasma sterilization (see below), can replace sterilization with ethylene oxide entirely. In addition, sterilization with ethylene oxide does not fail as frequently as sterilization with plasma when residual proteins and/or salts are present on the items (146).

Plasma Sterilization The low-temperature plasma is produced in a closed chamber with deep vacuum, an electromagnetic field, and a chemical precursor (hydrogen peroxide or a mixture of hydrogen peroxide and peracetic acid). The resulting free radicals, the chemical precursors, and the UV radiation are thought to be the products that rapidly destroy vegetative microorganisms including spores. The Sterrad 100 sterilizer was the first plasma sterilizer for use in health care facilities and has been on the market in Europe since 1990 and in the United States since 1993. In August 1997, the Sterrad 100 system was approved to sterilize certain surgical instruments with long lumens, such as those used in urologic, laparoscopic, and arthroscopic procedures, including instruments with single stainless steel lumens of ≥3 and 4 million single-use items without any published serious side effects, saving between 30 and 50% of the cost for a new item. With the expertise of an infection control professional, the health care facilities may provide the desired level of microbiological and toxicological safety. However, they probably cannot ensure that the design and function of the device are still adequate. Thus, in the United States and countries with similar regulations on quality assurance programs, reuse of single-use devices may not be cost-effective. In addition, organizations that sell used single-use devices to patients and/or insurance companies as new devices will encounter legal and ethical issues. Continuous improvements of new devices also impede reprocessing of singleuse devices on a large scale. However, financial restriction

206 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

may change the current beliefs; the reader should consult the FDA website and experts in the field before considering reprocessing single-use devices.

BSE AND vCJD CJD has been identified on all continents and is thought to occur worldwide. The incidence of CJD is estimated to be about 1 case per 106 persons per year. Most cases of CJD are sporadic; 10) followed by sterilization at 134°C for 5 min. Since 2002, Switzerland requires all surgical instruments to be sterilized at 134°C for 18 min. The background of the Swiss recommendation is that the usual rendering process for carcasses, which was discontinued, resulted in only a 1-log-unit reduction of the infectious particles (331). Therefore, a reduction in the number of infectious particles may suffice to stop transmission. CDC recommends that instruments exposed to potentially prion-contaminated items be autoclaved for 1 to 1.5 h at 132 to 134°C, immersed in 1 N sodium hydroxide for 1 h at room temperature, or immersed in sodium hypochlorite 0.5% (at least 2% free chlorine) for 2 h at room temperature.

13. Decontamination, Disinfection, and Sterilization n 207

See the CDC website for further information (http:// www.cdc.gov/ncidod/dvrd/vcjd/index.htm). In the United States, one patient who was a former resident of the United Kingdom has been diagnosed with vCJD, and the first case of BSE in cattle was identified in 2003. However, more cases may occur because 37 tons of “meals of meat or offal” that were “unfit for human consumption” was sent from the United Kingdom to the United States in 1997, well after the government banned imports of such risky meat (332). High-risk patients are patients with suspected CJD and their family members, patients treated with pituitary extracts, and patients who received cornea transplants. In addition, items should be considered contaminated with prions if a brain biopsy for the diagnosis of CJD is requested. Instruments used in such procedures should be discarded or placed under quarantine until the histopathological diagnosis is known. The incidence of vCJD in the United Kingdom is decreasing rapidly, indicating that current reprocessing techniques suffice. However, knowledge about this topic is increasing rapidly over time and our current understanding may be shown to be false in the future (254). In May 2005, British officials published an excellent assessment of the risk for contaminating surgical instruments with prions (https://www.gov.uk/ government/publications/guidance-from-the-acdp-tse-riskmanagement-subgroup-formerly-tse-working-group). The key observation in this report is that, on average, 0.2 mg of protein remains on surgical instruments despite “standard cleaning and disinfection,” which was sufficient to cause an experimental case of CJD. Therefore, more research and new methods of cleaning and disinfection are needed for surgical instruments. In laboratories, all samples should be handled using gloves and the processing of suspected specimens should be performed while wearing protective covering, e.g., gloves and disposable gowns or aprons (333). Masks and eyewear may be appropriate if splashing or aerosols are anticipated. Contaminations or spills should be decontaminated immediately with NaOH. To minimize environmental contamination, disposable cover sheets could be used on work surfaces (12). The reader is referred to the websites of the CDC, the FDA, and the WHO to obtain the most recent updates on this topic. In addition, the Society for Healthcare Epidemiology of America published detailed guidelines on this topic (12). More information about prions can be found in chapter 109 of this Manual. In conclusion, monitoring the processes of antiseptics, cleaning, disinfection, and sterilization is an essential part of infection control programs. The wide variety of applied techniques, germicidal agents, and procedures not only requires a broad understanding of the material, but it also needs well-trained personal to avoid incidents that result in damage to patients or equipment.

REFERENCES 1. Biron F, Verrier B, Peyramond D. 1997. Transmission of the human immunodeficiency virus and the hepatitis C virus. N Engl J Med 337:348–349. 2. Gillespie TG, Hogg L, Budge E, Duncan A, Coia JE. 2000. Mycobacterium chelonae isolated from rinse water within an endoscope washer-disinfector. J Hosp Infect 45:332–334. 3. Schelenz S, French G. 2000. An outbreak of multidrugresistant Pseudomonas aeruginosa infection associated with contamination of bronchoscopes and an endoscope washerdisinfector. J Hosp Infect 46:23–30.

4. Spach DH, Silverstein FE, Stamm WE. 1993. Transmission of infection by gastrointestinal endoscopy and bronchoscopy. Ann Intern Med 118:117–128. 5. Srinivasan A, Wolfenden LL, Song X, Mackie K, Hartsell TL, Jones HD, Diette GB, Orens JB, Yung RC, Ross TL, Merz W, Scheel PJ, Haponik EF, Perl TM. 2003. An outbreak of Pseudomonas aeruginosa infections associated with flexible bronchoscopes. N Engl J Med 348:221– 227. 6. Block SS. 2001. Disinfection, Sterilization, and Preservation. Lippincott Williams & Wilkins, Philadelphia, PA. 7. McDonnell G, Russell AD. 1999. Antiseptics and disinfectants: activity, action, and resistance. Clin Microbiol Rev 12:147–179. 8. Russell AD, Hugo WB, Ayliffe GAJ. 1992. Principles and Practice of Disinfection, Preservation and Sterilization. Blackwell Scientific Publications, London, United Kingdom. 9. Wenzel RP. 2004. The antibiotic pipeline—challenges, costs, and values. N Engl J Med 351:523–526. 10. Holmes KV. 2003. SARS-associated coronavirus. N Engl J Med 348:1948–1951. 11. Neumann G, Noda T, Kawaoka Y. 2009. Emergence and pandemic potential of swine-origin H1N1 influenza virus. Nature 459:931–939. 12. Rutala WA, Weber DJ. 2010. Guideline for disinfection and sterilization of prion-contaminated medical instruments. Infect Control Hosp Epidemiol 31:107–117. 13. Evans RJ. 1992. Acute anaphylaxis due to topical chlorhexidine acetate. BMJ 304:686. 14. Parker F, Foran S. 1995. Chlorhexidine catheter lubricant anaphylaxis. Anaesth Intensive Care 23:126. 15. Kramer A, Below H, Bieber N, Kampf G, Toma CD, Huebner NO, Assadian O. 2007. Quantity of ethanol absorption after excessive hand disinfection using three commercially available hand rubs is minimal and below toxic levels for humans. BMC Infect Dis 11:117. 16. Turner P, Saeed B, Kelsey MC. 2004. Dermal absorption of isopropyl alcohol from a commercial hand rub: implications for its use in hand decontamination. J Hosp Infect 56:287–290. 17. Rotter MA. 1996. Alcohols for antisepsis of hands and skin, p 177–234. In Ascenzi JP (ed), Handbook of Disinfectants and Antiseptics. Marcel Dekker, Inc, New York, NY. 18. Larson EL. 1991. Alcohols, p 191–203. In Block SS (ed), Disinfection, Sterilization and Preservation. Lea & Febiger, Philadelphia, PA. 19. Saurina G, Landman D, Quale JM. 1997. Activity of disinfectants against vancomycin-resistant Enterococcus faecium. Infect Control Hosp Epidemiol 18:345–347. 20. Rutala WA, et al. 1996. APIC guideline for selection and use of disinfectants. Am J Infect Control 24:313–342. 21. Klein M, DeForest A. 1963. The inactivation of viruses by germicides. Chem Specialists Manuf Assoc Proc 49:116– 118. 22. Mbithi JN, Springthorpe VS, Sattar SA. 1990. Chemical disinfection of hepatitis A virus on environmental surfaces. Appl Environ Microbiol 56:3601–3604. 23. Bond WW, Favero MS, Petersen NJ, Ebert JW. 1983. Inactivation of hepatitis B virus by intermediate-to-highlevel disinfectant chemicals. J Clin Microbiol 18:535–538. 24. Kobayashi H, Tsuzuki M, Koshimizu K, Toyama H, Yoshihara N, Shikata T, Abe K, Mizuno K, Otomo N, Oda T. 1984. Susceptibility of hepatitis B virus to disinfectants or heat. J Clin Microbiol 20:214–216. 25. Martin LS, McDougal JS, Loskoski SL. 1985. Disinfection and inactivation of the human T lymphotropic virus type III/lymphadenopathy-associated virus. J Infect Dis 152:400– 403. 26. van Bueren J, Larkin DP, Simpson RA. 1994. Inactivation of human immunodeficiency virus type 1 by alcohols. J Hosp Infect 28:137–148.

208 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

27. Kurtz JB, Lee TW, Parsons AJ. 1980. The action of alcohols on rotavirus, astrovirus and enterovirus. J Hosp Infect 1:321–325. 28. Ansari SA, Sattar SA, Springthorpe VS, Wells GA, Tostowaryk W. 1989. In vivo protocol for testing efficacy of hand-washing agents against viruses and bacteria: experiments with rotavirus and Escherichia coli. Appl Environ Microbiol 55:3113–3118. 29. Denton GE. 1991. Chlorhexidine, p 274–289. In Block SS (ed), Disinfection, Sterilization and Preservation. Lea & Febiger, Philadelphia, PA. 30. Ranganathan NS. 1996. Chlorhexidine, p 235–264. In Ascenzi JP (ed), Handbook of Disinfectants and Antiseptics. Marcel Dekker, Inc, New York, NY. 31. Maki DG, Stolz SM, Wheeler S, Mermel LA. 1997. Prevention of central venous catheter-related bloodstream infection by use of an antiseptic-impregnated catheter. A randomized, controlled trial. Ann Intern Med 127:257– 266. 32. Pronovost P, Needham D, Berenholtz S, Sinopoli D, Chu H, Cosgrove S, Sexton B, Hyzy R, Welsh R, Roth G, Bander J, Kepros J, Goeschel C. 2006. An intervention to decrease catheter-related bloodstream infections in the ICU. N Engl J Med 355:2725–2732. 33. Milstone AM, Passaretti CL, Perl TM. 2008. Chlorhexidine: expanding the armamentarium for infection control and prevention. Clin Infect Dis 46:274–281. 34. Larson EL, Butz AM, Gullette DL, Laughon BA. 1990. Alcohol for surgical scrubbing? Infect Control Hosp Epidemiol 11:139–143. 35. Dennis DT, Inglesby TV, Henderson DA, Bartlett JG, Ascher MS, Eitzen E, Fine AD, Friedlander AM, Hauer J, Layton M, Lillibridge SR, McDade JE, Osterholm MT, O’Toole T, Parker G, Perl TM, Russell PK, Tonat K. 2001. Tularemia as a biological weapon: medical and public health management. JAMA 285:2763–2773. 36. Lowbury EJ, Lilly HA. 1974. The effect of blood on disinfection of surgeons’ hands. Br J Surg 61:19–21. 37. Oie S, Kamiya A. 1996. Microbial contamination of antiseptics and disinfectants. Am J Infect Control 24:389–395. 38. Stickler DJ. 1974. Chlorhexidine resistance in Proteus mirabilis. J Clin Pathol 27:284–287. 39. Stickler DJ, Thomas B. 1980. Antiseptic and antibiotic resistance in Gram-negative bacteria causing urinary tract infection. J Clin Pathol 33:288–296. 40. Mayer S, Boos M, Beyer A, Fluit AC, Schmitz FJ. 2001. Distribution of the antiseptic resistance genes qacA, qacB and qacC in 497 methicillin-resistant and -susceptible European isolates of Staphylococcus aureus. J Antimicrob Chemother 47:896–897. 41. Mitchell BA, Brown MH, Skurray RA. 1998. QacA multidrug efflux pump from Staphylococcus aureus: comparative analysis of resistance to diamidines, biguanidines, and guanylhydrazones. Antimicrob Agents Chemother 42:475–477. 42. Yong D, Parker FC, Foran SM. 1995. Severe allergic reactions and intra-urethral chlorhexidine gluconate. Med J Aust 162:257–258. 43. Tabor E, Bostwick DC, Evans CC. 1989. Corneal damage due to eye contact with chlorhexidine gluconate. JAMA 261:557–558. 44. Marschall J, Mermel LA, et al. 2008. Strategies to prevent central line-associated bloodstream infections in acute care hospitals. Infect Control Hosp Epidemiol 29(Suppl 1):S22– S30. 45. Tamma PD, Aucott SW, Milstone AM. 2010. Chlorhexidine use in the neonatal intensive care unit: results from a national survey. Infect Control Hosp Epidemiol 31:846– 849. 46. Gottardi W. 1991. Iodine and iodine compounds, p 152– 166. In Block SS (ed), Disinfection, Sterilization and Preservation. Lea & Febiger, Philadelphia, PA.

47. Berkelman RL, Holland BW, Anderson RL. 1982. Increased bactericidal activity of dilute preparations of povidone-iodine solutions. J Clin Microbiol 15:635–639. 48. Berkelman RL, Lewin S, Allen JR, Anderson RL, Budnick LD, Shapiro S, Friedman SM, Nicholas P, Holzman RS, Haley RW. 1981. Pseudobacteremia attributed to contamination of povidone-iodine with Pseudomonas cepacia. Ann Intern Med 95:32–36. 49. Craven DE, Moody B, Connolly MG, Kollisch NR, Stottmeier KD, McCabe WR. 1981. Pseudobacteremia caused by povidone-iodine solution contaminated with Pseudomonas cepacia. N Engl J Med 305:621–623. 50. Anderson RL, Vess RW, Panlilio AL, Favero MS. 1990. Prolonged survival of Pseudomonas cepacia in commercially manufactured povidone-iodine. Appl Environ Microbiol 56:3598–3600. 51. Isenberg SJ, Apt L, Wood M. 1995. A controlled trial of povidone-iodine as prophylaxis against ophthalmia neonatorum. N Engl J Med 332:562–566. 52. Arata T, Murakami T, Hirai Y. 1993. Evaluation of povidone-iodine alcoholic solution for operative site disinfection. Postgrad Med J 69(Suppl 3):S93–S96. 53. Bryant WP, Zimmerman D. 1995. Iodine-induced hyperthyroidism in a newborn. Pediatrics 95:434–436. 54. Waran KD, Munsick RA. 1995. Anaphylaxis from povidone-iodine. Lancet 345:1506. 55. Jones RD, Jampani HB, Newman JL, Lee AS. 2000. Triclosan: a review of effectiveness and safety in health care settings. Am J Infect Control 28:184–196. 56. Suller MT, Russell AD. 2000. Triclosan and antibiotic resistance in Staphylococcus aureus. J Antimicrob Chemother 46:11–18. 57. Aiello AE, Larson E. 2003. Antibacterial cleaning and hygiene products as an emerging risk factor for antibiotic resistance in the community. Lancet Infect Dis 3:501–506. 58. Chuanchuen R, Beinlich K, Hoang TT, Becher A, Karkhoff-Schweizer RR, Schweizer HP. 2001. Cross-resistance between triclosan and antibiotics in Pseudomonas aeruginosa is mediated by multidrug efflux pumps: exposure of a susceptible mutant strain to triclosan selects nfxB mutants overexpressing MexCD-OprJ. Antimicrob Agents Chemother 45:428–432. 59. Russell AD. 2004. Whither triclosan? J Antimicrob Chemother 53:693–695. 60. Ghannoum MA, Elteen KA, Ellabib M, Whittaker PA. 1990. Antimycotic effects of octenidine and pirtenidine. J Antimicrob Chemother 25:237–245. 61. Sedlock DM, Bailey DM. 1985. Microbicidal activity of octenidine hydrochloride, a new alkanediylbis[pyridine] germicidal agent. Antimicrob Agents Chemother 28:786– 790. 62. Beiswanger BB, Mallatt ME, Mau MS, Jackson RD, Hennon DK. 1990. The clinical effects of a mouthrinse containing 0.1% octenidine. J Dent Res 69:454–457. 63. Tietz A, Frei R, Dangel M, Bolliger D, Passweg JR, Gratwohl A, Widmer AE. 2005. Octenidine hydrochloride for the care of central venous catheter insertion sites in severely immunocompromised patients. Infect Control Hosp Epidemiol 26:703–707. 64. Dettenkofer M, Wilson C, Gratwohl A, Schmoor C, Bertz H, Frei R, Heim D, Luft D, Schulz S, Widmer AF. 2010. Skin disinfection with octenidine dihydrochloride for central venous catheter site care: a double-blind, randomized, controlled trial. Clin Microbiol Infect 16:600–606. 65. Boyce JM, Pittet D. 2002. Guideline for hand hygiene in health-care settings: recommendations of the Healthcare Infection Control Practices Advisory Committee and the HICPAC/SHEA/APIC/IDSA Hand Hygiene Task Force. Infect Control Hosp Epidemiol 23:S3–S40. 66. Doebbeling BN, Stanley GL, Sheetz CT, Pfaller MA, Houston AK, Annis L, Li N, Wenzel RP. 1992. Compara-

13. Decontamination, Disinfection, and Sterilization n

67.

68.

69. 70.

71.

72.

73.

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

tive efficacy of alternative hand-washing agents in reducing nosocomial infections in intensive care units. N Engl J Med 327:88–93. Kampf G, Kramer A. 2004. Epidemiologic background of hand hygiene and evaluation of the most important agents for scrubs and rubs. Clin Microbiol Rev 17:863–893. Trampuz A, Widmer AF. 2004. Hand hygiene: a frequently missed lifesaving opportunity during patient care. Mayo Clin Proc 79:109–116. Widmer AF. 2000. Replace hand washing with use of a waterless alcohol hand rub? Clin Infect Dis 31:136–143. Price PB. 1938. The bacteriology of normal skin; a new quantitative test applied to a study of the bacterial biota and the disinfectant action of mechanical cleansing. J Infect Dis 63:301–318. Im SW, Fung JP, So SY, Yu DY. 1982. Unusual dissemination of pseudomonads by ventilators. Anaesthesia 37:1074– 1077. Adams BG, Marrie TJ. 1982. Hand carriage of aerobic Gram-negative rods may not be transient. J Hyg (London) 89:33–46. Boyce JM, Potter-Bynoe G, Opal SM, Dziobek L, Medeiros AA. 1990. A common-source outbreak of Staphylococcus epidermidis infections among patients undergoing cardiac surgery. J Infect Dis 161:493–499. Rutala WA, Weber DJ, Thomann CA, John JF, Saviteer SM, Sarubbi FA. 1988. An outbreak of Pseudomonas cepacia bacteremia associated with a contaminated intra-aortic balloon pump. J Thorac Cardiovasc Surg 96:157–161. Samore MH, Venkataraman L, DeGirolami PC, Arbeit RD, Karchmer AW. 1996. Clinical and molecular epidemiology of sporadic and clustered cases of nosocomial Clostridium difficile diarrhea. Am J Med 100:32–40. Widmer AF, Wenzel RP, Trilla A, Bale MJ, Jones RN, Doebbeling BN. 1993. Outbreak of Pseudomonas aeruginosa infections in a surgical intensive care unit: probable transmission via hands of a health care worker. Clin Infect Dis 16:372–376. Zaidi M, Sifuentes J, Bobadilla M, Moncada D, Ponce de León S. 1989. Epidemic of Serratia marcescens bacteremia and meningitis in a neonatal unit in Mexico City. Infect Control Hosp Epidemiol 10:14–20. Rotter ML, Simpson RA, Koller W. 1998. Surgical hand disinfection with alcohols at various concentrations: parallel experiments using the new proposed European standards method. Infect Control Hosp Epidemiol 19:778–781. Pittet D, Hugonnet S, Harbarth S, Mourouga P, Sauvan V, Touveneau S, Perneger TV. 2000. Effectiveness of a hospital-wide programme to improve compliance with hand hygiene. Lancet 356:1307–1312. Bellamy K, Alcock R, Babb JR, Davies JG, Ayliffe GA. 1993. A test for the assessment of ‘hygienic’ hand disinfection using rotavirus. J Hosp Infect 24:201–210. Rotter ML, Koller W, Wewalka G, Werner HP, Ayliffe GA, Babb JR. 1986. Evaluation of procedures for hygienic hand-disinfection: controlled parallel experiments on the Vienna test model. J Hyg (London) 96:27–37. Mermel LA, Josephson SL, Dempsey J, Parenteau S, Perry C, Magill N. 1997. Outbreak of Shigella sonnei in a clinical microbiology laboratory. J Clin Microbiol 35:3163– 3165. Katz KC, McGeer A, Low DE, Willey BM. 2002. Laboratory contamination of specimens with quality control strains of vancomycin-resistant enterococci in Ontario. J Clin Microbiol 40:2686–2688. Boyce JM, Pearson ML. 2003. Low frequency of fires from alcohol-based hand rub dispensers in healthcare facilities. Infect Control Hosp Epidemiol 24:618–619. Kramer A, Kampf G. 2007. Hand rub-associated fire incidents during 25,038 hospital-years in Germany. Infect Control Hosp Epidemiol 28:745–746.

209

86. Cruse PJ, Foord R. 1973. A five-year prospective study of 23,649 surgical wounds. Arch Surg 107:206–210. 87. Furuhashi M, Miyamae T. 1979. Effect of pre-operative hand scrubbing and influence of pinholes appearing in surgical rubber gloves during operation. Bull Tokyo Med Dent Univ 26:73–80. 88. Misteli H, Weber WP, Reck S, Rosenthal R, Zwahlen M, Fueglistaler P, Bolli MK, Oertli D, Widmer AF, Marti WR. 2009. Surgical glove perforation and the risk of surgical site infection. Arch Surg 144:553–558. 89. Fuursted K, Hjort A, Knudsen L. 1997. Evaluation of bactericidal activity and lag of regrowth (postantibiotic effect) of five antiseptics on nine bacterial pathogens. J Antimicrob Chemother 40:221–226. 90. Widmer AF. 2009. Surgical hand antisepsis, p 54–60. In World Health Organization (ed), WHO Guidelines on Hand Hygiene in Health Care. World Health Organization, Geneva, Switzerland. 91. Widmer AF, Rotter M, Voss A, Nthumba P, Allegranzi B, Boyce J, Pittet D. 2009. Surgical hand preparation: state-of-the-art. J Hosp Infect 74:112–122. 92. Blanc DS, Nahimana I, Petignat C, Wenger A, Bille J, Francioli P. 2004. Faucets as a reservoir of endemic Pseudomonas aeruginosa colonization/infections in intensive care units. Intensive Care Med 30:1964–1968. 93. Kramer A, Rudolph P, Kampf G, Pittet D. 2002. Limited efficacy of alcohol-based hand gels. Lancet 359:1489– 1490. 94. Kampf G, Ostermeyer C, Heeg P. 2005. Surgical hand disinfection with a propanol-based hand rub: equivalence of shorter application times. J Hosp Infect 59:304–310. 95. Weber WP, Reck S, Neff U, Saccilotto R, Dangel M, Rotter ML, Frei R, Oertli D, Marti WR, Widmer AF. 2009. Surgical hand antisepsis with alcohol-based hand rub: comparison of effectiveness after 1.5 and 3 minutes of application. Infect Control Hosp Epidemiol 30:420–426. 96. Garibaldi RA, Skolnick D, Lerer T, Poirot A, Graham J, Krisuinas E, Lyons R. 1988. The impact of preoperative skin disinfection on preventing intraoperative wound contamination. Infect Control Hosp Epidemiol 9:109–113. 97. Federal Register. 1994. Tentative final monograph for health-care antiseptic drug products. Fed Regist 59:31401– 31452. 98. Georgiade G, Riefkohl R, Georgiade N, Georgiade R, Wildman MF. 1985. Efficacy of povidone-iodine in preoperative skin preparation. J Hosp Infect 6(Suppl A):67– 71. 99. Swenson BR, Hedrick TL, Metzger R, Bonatti H, Pruett TL, Sawyer RG. 2009. Effects of preoperative skin preparation on postoperative wound infection rates: a prospective study of 3 skin preparation protocols. Infect Control Hosp Epidemiol 30:964–971. 100. Darouiche RO, Wall MJ Jr, Itani KM, Otterson MF, Webb AL, Carrick MM, Miller HJ, Awad SS, Crosby CT, Mosier MC, Alsharif A, Berger DH. 2010. Chlorhexidine-alcohol versus povidone-iodine for surgical-site antisepsis. N Engl J Med 362:18–26. 101. Tschudin-Sutter S, Frei R, Egli-Gany D, Eckstein F, Valderrabano V, Dangel M, Battegay M, Widmer AF. 2012. No risk of surgical site infections from residual bacteria after disinfection with povidone-iodine-alcohol in 1,014 cases: a prospective observational study. Ann Surg 255:565–569. 102. Dumville JC, McFarlane E, Edwards P, Lipp A, Holmes A. 2013. Preoperative skin antiseptics for preventing surgical wound infections after clean surgery. Cochrane Database Syst Rev 28:CD003949. 103. Mangram AJ, Horan TC, Pearson ML, Silver LC, Jarvis WR, et al. 1999. Guideline for prevention of surgical site infection, 1999. Infect Control Hosp Epidemiol 20:250– 278.

210 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

104. Webster J, Osborne S. 2012. Preoperative bathing or showering with skin antiseptics to prevent surgical site infection. Cochrane Database Syst Rev 12:CD004985. 105. Rotter ML, Larsen SO, Cooke EM, Dankert J, Daschner F, Greco D, Gronross P, Jepsen OB, Lystad A, Nystrom B, et al. 1988. A comparison of the effects of preoperative whole-body bathing with detergent alone and with detergent containing chlorhexidine gluconate on the frequency of wound infections after clean surgery. J Hosp Infect 11:310–320. 106. Lowbury EJ, Lilly HA, Bull JP. 1964. Methods for disinfection of operation sites. Br Med J 2:531–533. 107. Hill RL, Casewell MW. 2000. The in-vitro activity of povidone-iodine cream against Staphylococcus aureus and its bioavailability in nasal secretions. J Hosp Infect 45:198–205. 108. Kramer A, Roth B, Müller G, Rudolph P, Klöcker N. 2004. Influence of the antiseptic agents polyhexanide and octenidine on FL cells and on healing of experimental superficial aseptic wounds in piglets. A double-blind, randomised, stratified, controlled, parallel-group study. Skin Pharmacol Physiol 17:141–146. 109. Block C, Robenshtok E, Simhon A, Shapiro M. 2000. Evaluation of chlorhexidine and povidone iodine activity against methicillin-resistant Staphylococcus aureus and vancomycin-resistant Enterococcus faecalis using a surface test. J Hosp Infect 46:147–152. 110. Davies J, Babb JR, Ayliffe GA. 1977. The effect on the skin flora of bathing with antiseptic solutions. J Antimicrob Chemother 3:473–481. 111. Ayliffe GA, Noy MF, Babb JR, Davies JG, Jackson J. 1983. A comparison of pre-operative bathing with chlorhexidine-detergent and non-medicated soap in the prevention of wound infection. J Hosp Infect 4:237–244. 112. Wendt C, Schinke S, Württemberger M, Oberdorfer K, Bock-Hensley O, von Baum H. 2007. Value of wholebody washing with chlorhexidine for the eradication of methicillin-resistant Staphylococcus aureus: a randomized, placebo-controlled, double-blind clinical trial. Infect Control Hosp Epidemiol 28:1036–1043. 113. Leigh DA, Stronge JL, Marriner J, Sedgwick J. 1983. Total body bathing with ‘Hibiscrub’ (chlorhexidine) in surgical patients: a controlled trial. J Hosp Infect 4:229– 235. 114. Ridenour G, Lampen R, Federspiel J, Kritchevsky S, Wong E, Climo M. 2007. Selective use of intranasal mupirocin and chlorhexidine bathing and the incidence of methicillin-resistant Staphylococcus aureus colonization and infection among intensive care unit patients. Infect Control Hosp Epidemiol 28:1155–1161. 115. Hayek LJ, Emerson JM, Gardner AM. 1987. A placebocontrolled trial of the effect of two preoperative baths or showers with chlorhexidine detergent on postoperative wound infection rates. J Hosp Infect 10:165–172. 116. Cox RA, Conquest C. 1997. Strategies for the management of healthcare staff colonized with epidemic methicillin-resistant Staphylococcus aureus. J Hosp Infect 35:117– 127. 117. Valls V, Gómez-Herruz P, González-Palacios R, Cuadros JA, Romanyk JP, Ena J. 1994. Long-term efficacy of a program to control methicillin-resistant Staphylococcus aureus. Eur J Clin Microbiol Infect Dis 13:90–95. 118. Kotilainen P, Routamaa M, Peltonen R, Evesti P, Eerola E, Salmenlinna S, Vuopio-Varkila J, Rossi T. 2001. Eradication of methicillin-resistant Staphylococcus aureus from a health center ward and associated nursing home. Arch Intern Med 161:859–863. 119. Watanakunakorn C, Axelson C, Bota B, Stahl C. 1995. Mupirocin ointment with and without chlorhexidine baths in the eradication of Staphylococcus aureus nasal

120.

121.

122.

123.

124.

125.

126.

127.

128.

129.

130.

131.

132.

133.

carriage in nursing home residents. Am J Infect Control 23:306–309. Sandri AM, Dalarosa MG, Ruschel de Alcantara L, da Silva Elias L, Zavascki AP. 2006. Reduction in incidence of nosocomial methicillin-resistant Staphylococcus aureus (MRSA) infection in an intensive care unit: role of treatment with mupirocin ointment and chlorhexidine baths for nasal carriers of MRSA. Infect Control Hosp Epidemiol 27:185–187. Huang SS, Septimus E, Kleinman K, Moody J, Hickok J, Avery TR, Lankiewicz J, Gombosev A, Terpstra L, Hartford F, Hayden MK, Jernigan JA, Weinstein RA, Fraser VJ, Haffenreffer K, Cui E, Kaganov RE, Lolans K, Perlin JB, Platt R, CDC Prevention Epicenters Program, AHRQ DECIDE Network and Healthcare-Associated Infections Program. 2013. Targeted versus universal decolonization to prevent ICU infection. N Engl J Med 368:2255–2265. Rohr U, Mueller C, Wilhelm M, Muhr G, Gatermann S. 2003. Methicillin-resistant Staphylococcus aureus wholebody decolonization among hospitalized patients with variable site colonization by using mupirocin in combination with octenidine dihydrochloride. J Hosp Infect 54:305–309. Sloot N, Siebert J, Hoffler U. 1999. Eradication of MRSA from carriers by means of whole-body washing with an antiseptic in combination with mupirocin nasal ointment. Zentralbl Hyg Umweltmed 202:513–523. Cohen SH, Morita MM, Bradford M. 1991. A sevenyear experience with methicillin-resistant Staphylococcus aureus. Am J Med 91:233S–237S. Zafar AB, Butler RC, Reese DJ, Gaydos LA, Mennonna PA. 1995. Use of 0.3% triclosan (Bacti-Stat) to eradicate an outbreak of methicillin-resistant Staphylococcus aureus in a neonatal nursery. Am J Infect Control 23:200–208. Nguyen DM, Mascola L, Brancoft E. 2005. Recurring methicillin-resistant Staphylococcus aureus infections in a football team. Emerg Infect Dis 11:526–532. Wilcox MH, Hall J, Gill AB, Fawley WN, Parnell P, Verity P. 2004. Effectiveness of topical chlorhexidine powder as an alternative to hexachlorophane for the control of Staphylococcus aureus in neonates. J Hosp Infect 56:156–159. Kampf G, Kramer A. 2004. Eradication of methicillinresistant Staphylococcus aureus with an antiseptic soap and nasal mupirocin among colonized patients—an open uncontrolled clinical trial. Ann Clin Microbiol Antimicrob 3:9. Fukunaga A, Naritaka H, Fukaya R, Tabuse M, Nakamura T. 2004. Povidone-iodine ointment and gauze dressings associated with reduced catheter-related infection in seriously ill neurosurgical patients. Infect Control Hosp Epidemiol 25:696–698. Reimer K, Wichelhaus TA, Schäfer V, Rudolph P, Kramer A, Wutzler P, Ganzer D, Fleischer W. 2002. Antimicrobial effectiveness of povidone-iodine and consequences for new application areas. Dermatology 204(Suppl 1):114–120. Seaman PF, Ochs D, Day MJ. 2007. Small-colony variants: a novel mechanism for triclosan resistance in methicillin-resistant Staphylococcus aureus. J Antimicrob Chemother 59:43–50. Bayston R, Ashraf W, Smith T. 2007. Triclosan resistance in methicillin-resistant Staphylococcus aureus expressed as small colony variants: a novel mode of evasion of susceptibility to antiseptics—authors’ response. J Antimicrob Chemother 60:176–177. Brenwald NP, Fraise AP. 2003. Triclosan resistance in methicillin-resistant Staphylococcus aureus (MRSA). J Hosp Infect 55:141–144

13. Decontamination, Disinfection, and Sterilization n 134. Ringberg H, Petersson AC, Walder M, Johansson PJH. 2006. The throat: an important site for MRSA colonization. Scand J Infect Dis 38:888–893. 135. Brook I, Foote PA. 2006. Isolation of methicillin resistant Staphylococcus aureus from the surface and core of tonsils in children. Int J Pediatr Otorhinolaryngol 70:2099–2102. 136. Pitten FA, Werner HP, Kramer A. 2003. A standardized test to assess the impact of different organic challenges on the antimicrobial activity of antiseptics. J Hosp Infect 55:108–115. 137. Tomás I, Alvarez M, Limeres J, Tomás M, Medina J, Otero JL, Diz P. 2007. Effect of a chlorhexidine mouthwash on the risk of postextraction bacteremia. Infect Control Hosp Epidemiol 28:577–582. 138. Pitten FA, Kiefer T, Buth C, Doelken G, Kramer A. 2003. Do cancer patients with chemotherapy-induced leukopenia benefit from an antiseptic chlorhexidine-based oral rinse? A double-blind, block-randomized, controlled study. J Hosp Infect 53:283–291. 139. van Steenberghe D. 1997. Breath malodor. Curr Opin Periodontol 4:137–143. 140. Clavero J, Baca P, Junco P, González MP. 2003. Effects of 0.2% chlorhexidine spray applied once or twice daily on plaque accumulation and gingival inflammation in a geriatric population. J Clin Periodontol 30:773–777. 141. Leenders AC, Renders NR, Pelk M, Janssen M. 2005. Tonsillectomy for treatment of persistent methicillin-resistant Staphylococcus aureus throat carriage. J Hosp Infect 59:266–267. 142. Patterson P. 2009. CDC sterilization, disinfection guideline. OR Manager 25:14–16. 143. Fichet G, Comoy E, Duval C, Antloga K, Dehen C, Charbonnier A, McDonnell G, Brown P, Lasmezas CI, Deslys JP. 2004. Novel methods for disinfection of prioncontaminated medical devices. Lancet 364:521–526. 144. Hanson PJ, Jeffries DJ, Collins JV. 1991. Viral transmission and fibreoptic endoscopy. J Hosp Infect 18(Suppl A):136–140. 145. Nelson DB, Jarvis WR, Rutala WA, Foxx-Orenstein AE, Isenberg G, Dash GR, Alvarado CJ, Ball M, GriffinSobel J, Petersen C, Ball KA, Henderson J, Stricof RL, et al. 2003. Multi-society guideline for reprocessing flexible gastrointestinal endoscopes. Infect Control Hosp Epidemiol 24:532–537. 146. Alfa MJ, DeGagne P, Olson N. 1997. Bacterial killing ability of 10% ethylene oxide plus 90% hydrochlorofluorocarbon sterilizing gas. Infect Control Hosp Epidemiol 18:641–645. 147. Dharan S, Mourouga P, Copin P, Bessmer G, Tschanz B, Pittet D. 1999. Routine disinfection of patients’ environmental surfaces. Myth or reality? J Hosp Infect 42:113– 117. 148. Sehulster L, Chinn RY. 2003. Guidelines for environmental infection control in health-care facilities. Recommendations of CDC and the Healthcare Infection Control Practices Advisory Committee (HICPAC). MMWR Recommend. Rep. 52:1–42. 149. Weber DJ, Rutala WA, Sickbert-Bennett EE. 2007. Outbreaks associated with contaminated antiseptics and disinfectants. Antimicrob Agents Chemother 51:4217–4224. 150. Reiss I, Borkhardt A, Fussle R, Sziegoleit A, Gortner L. 2000. Disinfectant contaminated with Klebsiella oxytoca as a source of sepsis in babies. Lancet 356:310. 151. Astagneau P, Desplaces N, Vincent V, Chicheportiche V, Botherel A, Maugat S, Lebascle K, Leonard P, Desenclos J, Grosset J, Ziza J, Brucker G. 2001. Mycobacterium xenopi spinal infections after discovertebral surgery: investigation and screening of a large outbreak. Lancet 358:747–751.

211

152. Tyler R, Ayliffe GA, Bradley C. 1990. Virucidal activity of disinfectants: studies with the poliovirus. J Hosp Infect 15:339–345. 153. Rutala WA, Weber DJ. 1995. FDA labeling requirements for disinfection of endoscopes: a counterpoint. Infect Control Hosp Epidemiol 16:231–235. 154. Hanson PJ, Gor D, Clarke JR, Chadwick MV, Gazzard B, Jeffries DJ, Gaya H, Collins JV. 1991. Recovery of the human immunodeficiency virus from fibreoptic bronchoscopes. Thorax 46:410–412. 155. Ascenzi JM. 1996. Glutaraldehyde-based disinfectants, p 111–132. In Ascenzi JP (ed), Handbook of Disinfectants and Antiseptics. Marcel Dekker, Inc, New York, NY. 156. Russell AD. 1994. Glutaraldehyde: current status and uses. Infect Control Hosp Epidemiol 15:724–733. 157. Gelinas P, Goulet J, Tastayre GM, Picard GA. 1991. Effect of temperature and contact time on the activity of eight disinfectants—a classification. J Food Prot 47:841– 847. 158. Rutala WA, Gergen MF, Weber DJ. 1993. Inactivation of Clostridium difficile spores by disinfectants. Infect Control Hosp Epidemiol 14:36–39. 159. Wilson JA, Margolin AB. 1999. The efficacy of three common hospital liquid germicides to inactivate Cryptosporidium parvum oocysts. J Hosp Infect 42:231–237. 160. Rubbo SD, Gardner JF, Webb RL. 1967. Biocidal activities of glutaraldehyde and related compounds. J Appl Bacteriol 30:78–87. 161. Collins FM. 1986. Kinetics of the tuberculocidal response by alkaline glutaraldehyde in solution and on an inert surface. J Appl Bacteriol 61:87–93. 162. Collins FM. 1986. Bactericidal activity of alkaline glutaraldehyde solution against a number of atypical mycobacterial species. J Appl Bacteriol 61:247–251. 163. Jackson J, Leggett JE, Wilson DA, Gilbert DN. 1996. Mycobacterium gordonae in fiberoptic bronchoscopes. Am J Infect Control 24:19–23. 164. Griffiths PA, Babb JR, Bradley CR, Fraise AP. 1997. Glutaraldehyde-resistant Mycobacterium chelonae from endoscope washer disinfectors. J Appl Microbiol 82:519–526. 165. Van Klingeren B, Pullen W. 1993. Glutaraldehyde resistant mycobacteria from endoscope washers. J Hosp Infect 25:147–149. 166. Mbithi JN, Springthorpe VS, Sattar SA, Pacquette M. 1993. Bactericidal, virucidal, and mycobactericidal activities of reused alkaline glutaraldehyde in an endoscopy unit. J Clin Microbiol 31:2988–2995. 167. Alvarado CJ, Reichelderfer M, et al. 2000. APIC guideline for infection prevention and control in flexible endoscopy. Am J Infect Control 28:138–155. 168. Dolce P, Gourdeau M, April N, Bernard PM. 1995. Outbreak of glutaraldehyde-induced proctocolitis. Am J Infect Control 23:34–39. 169. West AB, Kuan SF, Bennick M, Lagarde S. 1995. Glutaraldehyde colitis following endoscopy: clinical and pathological features and investigation of an outbreak. Gastroenterology 108:1250–1255. 170. Brown P, Liberski PP, Wolff A, Gajdusek DC. 1990. Resistance of scrapie infectivity to steam autoclaving after formaldehyde fixation and limited survival after ashing at 360 degrees C: practical and theoretical implications. J Infect Dis 161:467–472. 171. Fraud S, Maillard JY, Russell AD. 2001. Comparison of the mycobactericidal activity of ortho-phthalaldehyde, glutaraldehyde and other dialdehydes by a quantitative suspension test. J Hosp Infect 48:214–221. 172. Walsh SE, Maillard JY, Russell AD. 1999. Ortho-phthalaldehyde: a possible alternative to glutaraldehyde for high level disinfection. J Appl Microbiol 86:1039–1046.

212 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

173. Rossmoore HW, Sondossi M. 1988. Applications and mode of action of formaldehyde condensate biocides. Adv Appl Microbiol 33:223–277. 174. Akbar-Khanzadeh F, Vaquerano MU, Akbar-Khanzadeh M, Bisesi MS. 1994. Formaldehyde exposure, acute pulmonary response, and exposure control options in a gross anatomy laboratory. Am J Ind Med 26:61–75. 175. Rutala WA, Weber DJ. 1997. Uses of inorganic hypochlorite (bleach) in health-care facilities. Clin Microbiol Rev 10:597–610. 176. Karol MH. 1995. Toxicologic principles do not support the banning of chlorine. A Society of Toxicology position paper. Fundam Appl Toxicol 24:1–2. 177. Williams ND, Russell AD. 1991. The effects of some halogen-containing compounds on Bacillus subtilis endospores. J Appl Bacteriol 70:427–436. 178. Kaatz GW, Gitlin SD, Schaberg DR, Wilson KH, Kauffman CA, Seo SM, Fekety R. 1988. Acquisition of Clostridium difficile from the hospital environment. Am J Epidemiol 127:1289–1293. 179. Centers for Disease Control and Prevention. 2001. Protracted outbreaks of cryptosporidiosis associated with swimming pool use—Ohio and Nebraska, 2000. MMWR Morb Mortal Wkly Rep 50:406–410. 180. Helms CM, Massanari RM, Wenzel RP, Pfaller MA, Moyer NP, Hall N. 1988. Legionnaires’ disease associated with a hospital water system. A five-year progress report on continuous hyperchlorination. JAMA 259:2423–2427. 181. Centers for Disease Control and Prevention. 1989. Guidelines for prevention of transmission of human immunodeficiency virus and hepatitis B virus to health-care and public-safety workers. MMWR Morb Mortal Wkly Rep 38(Suppl 6):1–37. 182. Chitnis V, Chitnis S, Patil S, Chitnis D. 2004. Practical limitations of disinfection of body fluid spills with 10,000 ppm sodium hypochlorite (NaOCl). Am J Infect Control 32:306–308. 183. Flynn N, Jain S, Keddie EM, Carlson JR, Jennings MB, Haverkos HW, Nassar N, Anderson R, Cohen S, Goldberg D. 1994. In vitro activity of readily available household materials against HIV-1: is bleach enough? J Acquir Immune Defic Syndr 7:747–753. 184. Weber DJ, Barbee SL, Sobsey MD, Rutala WA. 1999. The effect of blood on the antiviral activity of sodium hypochlorite, a phenolic, and a quaternary ammonium compound. Infect Control Hosp Epidemiol 20:821–827. 185. Centers for Disease Control and Prevention. 1996. Community-level prevention of human immunodeficiency virus infection among high-risk populations: the AIDS community demonstration projects. MMWR Recommend Rep 45(RR-6):1–31. 186. Hamilton E, Seal DV, Hay J. 1996. Comparison of chlorine and chlorine dioxide disinfection for control of Legionella in a hospital potable water supply. J Hosp Infect 32:156–160. 187. Kool JL, Carpenter JC, Fields BS. 1999. Effect of monochloramine disinfection of municipal drinking water on risk of nosocomial Legionnaires’ disease. Lancet 353:272– 277. 188. Lever AM, Sutton SVW. 1996. Antimicrobial effects of hydrogen peroxide as an antiseptic and disinfectant, p 159–176. In Ascenzi JP (ed), Handbook of Disinfectants and Antiseptics. Marcel Dekker, Inc, New York, NY. 189. Mathers WD, Sutphin JE, Folberg R, Meier PA, Wenzel RP, Elgin RG. 1996. Outbreak of keratitis presumed to be caused by Acanthamoeba. Am J Ophthalmol 121:129– 142. 190. Wardle MD, Renninger GM. 1975. Bactericidal effect of hydrogen peroxide on spacecraft isolates. Appl Microbiol 30:710–711.

191. Leaper S. 1984. Influence of temperature on the synergistic sporicidal effect of peracetic acid plus hydrogen peroxide in Bacillus subtilis SA22 (NCA 72–52). Food Microbiol 1:199–203. 192. Sagripanti JL, Bonifacino A. 1996. Comparative sporicidal effect of liquid chemical germicides on three medical devices contaminated with spores of Bacillus subtilis. Am J Infect Control 24:364–371. 193. Alasri A, Valverde M, Roques C, Michel G, Cabassud C, Aptel P. 1993. Sporocidal properties of peracetic acid and hydrogen peroxide, alone and in combination, in comparison with chlorine and formaldehyde for ultrafiltration membrane disinfection. Can J Microbiol 39:52– 60. 194. Mentel R, Schmidt J. 1973. Investigations on rhinovirus inactivation by hydrogen peroxide. Acta Virol 17:351– 354. 195. Ryan CK, Potter GD. 1995. Disinfectant colitis. Rinse as well as you wash. J Clin Gastroenterol 21:6–9. 196. Hughes R, Kilvington S. 2001. Comparison of hydrogen peroxide contact lens disinfection systems and solutions against Acanthamoeba polyphaga. Antimicrob Agents Chemother 45:2038–2043. 197. Levenson JE. 1989. Corneal damage from improperly cleaned tonometer tips. Arch Ophthalmol 107:1117. 198. Alasri A, Roques C, Michel G, Cabassud C, Aptel P. 1992. Bactericidal properties of peracetic acid and hydrogen peroxide, alone and in combination, and chlorine and formaldehyde against bacterial water strains. Can J Microbiol 38:635–642. 199. Sagripanti JL, Eklund CA, Trost PA, Jinneman KC, Abeyta CJ, Kaysner CA, Hill WE. 1997. Comparative sensitivity of 13 species of pathogenic bacteria to seven chemical germicides. Am J Infect Control 25:335–339. 200. Held PJ, Wolfe RA, Gaylin DS, Port FK, Levin NW, Turenne MN. 1994. Analysis of the association of dialyzer reuse practices and patient outcomes. Am J Kidney Dis 23:692–708. 201. Feldman HI, Kinosian M, Bilker WB, Simmons C, Holmes JH, Pauly MV, Escarce JJ. 1996. Effect of dialyzer reuse on survival of patients treated with hemodialysis. JAMA 276:1724. 202. O’Connor DO, Rubino JR. 1991. Phenolic compounds, p 204–224. In Block SS (ed), Disinfection, Sterilization and Preservation. Lea & Febiger, Philadelphia, PA. 203. Rutala WA, Cole EC. 1987. Ineffectiveness of hospital disinfectants against bacteria: a collaborative study. Infect Control 8:501–506. 204. Terleckyj B, Axler DA. 1987. Quantitative neutralization assay of fungicidal activity of disinfectants. Antimicrob Agents Chemother 31:794–798. 205. Narang HK, Codd AA. 1983. Action of commonly used disinfectants against enteroviruses. J Hosp Infect 4:209– 212. 206. Druce JD, Jardine D, Locarnini SA, Birch CJ. 1995. Susceptibility of HIV to inactivation by disinfectants and ultraviolet light. J Hosp Infect 30:167–180. 207. Sattar SA, Jacobsen H, Rahman H, Cusack TM, Rubino JR. 1994. Interruption of rotavirus spread through chemical disinfection. Infect Control Hosp Epidemiol 15:751– 756. 208. Merianos JJ. 1991. Quaternary ammonium antimicrobial compounds, p 225–255. In Block SS (ed), Disinfection, Sterilization and Preservation. Lea & Febiger, Philadelphia, PA. 209. Rutala WA, Cole EC, Wannamaker NS, Weber DJ. 1991. Inactivation of Mycobacterium tuberculosis and Mycobacterium bovis by 14 hospital disinfectants. Am J Med 91:267S–271S.

13. Decontamination, Disinfection, and Sterilization n 210. Frank MJ, Schaffner W. 1976. Contaminated aqueous benzalkonium chloride. An unnecessary hospital infection hazard. JAMA 236:2418–2419. 211. Nakashima AK, Highsmith AK, Martone WJ. 1987. Survival of Serratia marcescens in benzalkonium chloride and in multiple-dose medication vials: relationship to epidemic septic arthritis. J Clin Microbiol 25:1019–1021. 212. Tiwari TS, Ray B, Jost KC Jr, Rathod MK, Zhang Y, Brown-Elliott BA, Hendricks K, Wallace RJ Jr. 2003. Forty years of disinfectant failure: outbreak of postinjection Mycobacterium abscessus infection caused by contamination of benzalkonium chloride. Clin Infect Dis 36:954– 962. 213. Casalegno JS, Le Bail Carval K, Eibach D, Valdeyron ML, Lamblin G, Jacquemoud H, Mellier G, Lina B, Gaucherand P, Mathevet P, Mekki Y. 2012. High risk HPV contamination of endocavity vaginal ultrasound probes: an underestimated route of nosocomial infection? PLoS One 7:e48137. 214. Disch K. 1994. Glucoprotamine—a new antimicrobial substance. Zentralbl Hyg Umweltmed 195:357–365. 215. Meyer B, Kluin C. 1999. Efficacy of Glucoprotamin containing disinfectants against different species of atypical mycobacteria. J Hosp Infect 42:151–154. 216. Widmer AF. 2003. Antimicrobial activity of glucoprotamin: a clinical study of a new disinfectant for instruments. Infect Control Hosp Epidemiol 24:762–764. 217. Coates D. 1996. Sporicidal activity of sodium dichloroisocyanurate, peroxygen and glutaraldehyde disinfectants against Bacillus subtilis. J Hosp Infect 32:283–294. 218. Broadley SJ, Furr JR, Jenkins PA, Russell AD. 1993. Antimycobacterial activity of ‘Virkon’. J Hosp Infect 23:189–197. 219. Middleton AM, Chadwick MV, Sanderson JL, Gaya H. 2000. Comparison of a solution of super-oxidized water (Sterilox) with glutaraldehyde for the disinfection of bronchoscopes, contaminated. J Hosp Infect 45:278–282. 220. Shetty N, Srinivasan S, Holton J, Ridgway GL. 1999. Evaluation of microbicidal activity of a new disinfectant: Sterilox 2500 against Clostridium difficile spores, Helicobacter pylori, vancomycin resistant Enterococcus species, Candida albicans and several Mycobacterium species. J Hosp Infect 41:101–105. 221. Sagripanti JL. 1992. Metal-based formulations with high microbicidal activity. Appl Environ Microbiol 58:3157– 3162. 222. Stout JE, Lin YS, Goetz AM, Muder RR. 1998. Controlling Legionella in hospital water systems: experience with the superheat-and-flush method and copper-silver ionization. Infect Control Hosp Epidemiol 19:911–914. 223. Brady MJ, Lisay CM, Yurkovetskiy AV, Sawan SP. 2003. Persistent silver disinfectant for the environmental control of pathogenic bacteria. Am J Infect Control 31:208–214. 224. Rutala WA, Weber DJ. 2001. New disinfection and sterilization methods. Emerg Infect Dis 7:348–353. 225. Archibald LK, Corl A, Shah B, Schulte M, Arduino MJ, Aguero S, Fisher DJ, Stechenberg BW, Banerjee SN, Jarvis WR. 1997. Serratia marcescens outbreak associated with extrinsic contamination of 1% chlorxylenol soap. Infect Control Hosp Epidemiol 18:704–709. 226. Bosi C, Davin-Regli A, Charrel R, Rocca B, Monnet D, Bollet C. 1996. Serratia marcescens nosocomial outbreak due to contamination of hexetidine solution. J Hosp Infect 33:217–224. 227. Vigeant P, Loo VG, Bertrand C, Dixon C, Hollis R, Pfaller MA, McLean AP, Briedis DJ, Perl TM, Robson HG. 1998. An outbreak of Serratia marcescens infections related to contaminated chlorhexidine. Infect Control Hosp Epidemiol 19:791–794.

213

228. Tschudin-Sutter S, Frei R, Kampf G, Tamm M, Pflimlin E, Battegay M, Widmer AF. 2011. Emergence of glutaraldehyde-resistant Pseudomonas aeruginosa. Infect Control Hosp Epidemiol 32:1173–1178. 229. Perencevich EN, Wong MT, Harris AD. 2001. National and regional assessment of the antibacterial soap market: a step toward determining the impact of prevalent antibacterial soaps. Am J Infect Control 29:281–283. 230. Fraise AP, Lambert PA, Maillard JY. 2004. Russell, Hugo and Ayliffe’s Principles and Practice of Disinfections, Preservation and Sterilization. Blackwell Publishing, Malden, MA. 231. Maillard JY. 2002. Bacterial target sites for biocide action. J Appl Microbiol 92(Suppl):16S–27S. 232. Sheldon AT Jr. 2005. Antiseptic “resistance”: real or perceived threat? Clin Infect Dis 40:1650–1656. 233. Weber DJ, Sickbert-Bennett E, Gergen MF, Rutala WA. 2003. Efficacy of selected hand hygiene agents used to remove Bacillus atrophaeus (a surrogate of Bacillus anthracis) from contaminated hands. JAMA 289:1274–1277. 234. Rutala WA, Weber DJ. 2004. Disinfection and sterilization in health care facilities: what clinicians need to know. Clin Infect Dis 39:702–709. 235. Lai MY, Cheng PK, Lim WW. 2005. Survival of severe acute respiratory syndrome coronavirus. Clin Infect Dis 41:e67–e71. 236. Wullt M, Odenholt I, Walder M. 2003. Activity of three disinfectants and acidified nitrite against Clostridium difficile spores. Infect Control Hosp Epidemiol 24:765–768. 237. Orenstein R, Aronhalt KC, McManus JE Jr, Fedraw LA. 2011. A targeted strategy to wipe out Clostridium difficile. Infect Control Hosp Epidemiol 32:1137–1139. 238. Fenner L, Widmer AF, Stranden A, Conzelmann M, Goorhuis A, Harmanus C, Kuijper EJ, Frei R. 2008. First cluster of clindamycin-resistant Clostridium difficile PCR ribotype 027 in Switzerland. Clin Microbiol Infect 14:514–515. 239. McDonald LC, Killgore GE, Thompson A, Owens RC Jr, Kazakova SV, Sambol SP, Johnson S, Gerding DN. 2005. An epidemic, toxin gene-variant strain of Clostridium difficile. N Engl J Med 353:2433–2441. 240. Wilcox MH, Fawley WN. 2000. Hospital disinfectants and spore formation by Clostridium difficile. Lancet 356:1324. 241. Hall CB, Douglas RG Jr. 1981. Modes of transmission of respiratory syncytial virus. J Pediatr 99:100–103. 242. Gamage B, Moore D, Copes R, Yassi A, Bryce E. 2005. Protecting HCWs from SARS and other respiratory pathogens: a review of the infection control literature. Am J Infect Control 33:114–121. 243. Wysowski DK, Flynt JWJ, Goldfield M, Altman R, Davis AT. 1978. Epidemic neonatal hyperbilirubinemia and use of a phenolic disinfectant detergent. Pediatrics 61:165–170. 244. de Lassence A, Hidri N, Timsit JF, Joly-Guillou ML, Thiery G, Boyer A, Lable P, Blivet A, Kalinowski H, Martin Y, Lajonchere JP, Dreyfuss D. 2006. Control and outcome of a large outbreak of colonization and infection with glycopeptide-intermediate Staphylococcus aureus in an intensive care unit. Clin Infect Dis 42:170–178. 245. Clay S, Maherchandani S, Malik YS, Goyal SM. 2006. Survival on uncommon fomites of feline calicivirus, a surrogate of noroviruses. Am J Infect Control 34:41–43. 246. Dancer SJ. 2008. Importance of the environment in methicillin-resistant Staphylococcus aureus acquisition: the case for hospital cleaning. Lancet Infect Dis 8:101–113. 247. Khanna N, Goldenberger D, Graber P, Battegay M, Widmer AF. 2003. Gastroenteritis outbreak with norovirus in a Swiss university hospital with a newly identified virus strain. J Hosp Infect 55:131–136.

214 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

248. Boyce JM, Havill NL, Otter JA, McDonald LC, Adams NM, Cooper T, Thompson A, Wiggs L, Killgore G, Tauman A, Noble-Wang J. 2008. Impact of hydrogen peroxide vapor room decontamination on Clostridium difficile environmental contamination and transmission in a healthcare setting. Infect Control Hosp Epidemiol 29:723–729. 249. Passaretti CL, Otter JA, Reich NG, Myers J, Shepard J, Ross T, Carroll KC, Lipsett P, Perl TM. 2013. An evaluation of environmental decontamination with hydrogen peroxide vapor for reducing the risk of patient acquisition of multidrug-resistant organisms. Clin Infect Dis 56:27–35. 250. Jeanes A, Rao G, Osman M, Merrick P. 2005. Eradication of persistent environmental MRSA. J Hosp Infect 61:85–86 251. Dryden M, Parnaby R, Dailly S, Lewis T, Davis-Blues K, Otter JA, Kearns AM. 2008. Hydrogen peroxide vapour decontamination in the control of a polyclonal methicillin-resistant Staphylococcus aureus outbreak on a surgical ward. J Hosp Infect 68:190–192. 252. Otter JA, Yezli S, Schouten MA, van Zanten AR, Houmes-Zielman G, Nohlmans-Paulssen MK. 2010 Hydrogen peroxide vapor decontamination of an intensive care unit to remove environmental reservoirs of multidrug-resistant Gram-negative rods during an outbreak. Am J Infect Control 38:754–756 253. Otter JA, Yezli S, Perl MZ, Barbut F, French GL. 2013. The role of ‘no-touch’ automated room disinfection systems in infection prevention and control. J Hosp Infect 83:1–13. 254. Belay ED, Schonberger LB. 2005. The public health impact of prion diseases. Annu Rev Public Health 26:191– 212. 255. Collins SJ, Lawson VA, Masters CL. 2004. Transmissible spongiform encephalopathies. Lancet 363:51–61. 256. Weissmann C. 2005. Birth of a prion: spontaneous generation revisited. Cell 122:165–168. 257. Spaulding EH. 1968. Chemical disinfection of medical and surgical materials, p 517–531. In Block S (ed), Disinfection, Sterilization and Preservation. Lea & Febiger, Philadelphia, PA. 258. Prusiner SB. 1982. Novel proteinaceous infectious particles cause scrapie. Science 216:136–144. 259. Cetinkaya Y, Falk P, Mayhall CG. 2000. Vancomycinresistant enterococci. Clin Microbiol Rev 13:686–707. 260. Martin MA, Reichelderfer M, et al. 1994. APIC guidelines for infection prevention and control in flexible endoscopy. Am J Infect Control 22:19–38. 261. Corne P, Godreuil S, Jean-Pierre H, Jonquet O, Campos J, Jumas-Bilak E, Parer S, Marchandin H. 2005. Unusual implication of biopsy forceps in outbreaks of Pseudomonas aeruginosa infections and pseudo-infections related to bronchoscopy. J Hosp Infect 61:20–26. 262. Deva AK, Vickery K, Zou J, West RH, Harris JP, Cossart YE. 1996. Establishment of an in-use testing method for evaluating disinfection of surgical instruments using the duck hepatitis B model. J Hosp Infect 33:119– 130. 263. Wu MS, Wang JT, Yang JC, Wang HH, Sheu JC, Chen DS, Wang TH. 1996. Effective reduction of Helicobacter pylori infection after upper gastrointestinal endoscopy by mechanical washing of the endoscope. Hepatogastroenterology 43:1660–1664. 264. Kaczmarek RG, Moore RMJ, McCrohan J, Goldmann DA, Reynolds C, Caquelin C, Israel E. 1992. Multistate investigation of the actual disinfection/sterilization of endoscopes in health care facilities. Am J Med 92:257– 261.

265. Favero MS. 1991. Strategies for disinfection and sterilization of endoscopes: the gap between basic principles and actual practice. Infect Control Hosp Epidemiol 12:279–281. 266. Gorse GJ, Messner RL. 1991. Infection control practices in gastrointestinal endoscopy in the United States: a national survey. Infect Control Hosp Epidemiol 12:289–296. 267. Cheung RJ, Ortiz D, DiMarino AJ Jr. 1999. GI endoscopic reprocessing practices in the United States. Gastrointest Endosc 50:362–368. 268. Robertson LJ, Campbell AT, Smith HV. 1992. Survival of Cryptosporidium parvum oocysts under various environmental pressures. Appl Environ Microbiol 58:3494–3500. 269. Akamatsu T, Tabata K, Hironaga M, Uyeda M. 1997. Evaluation of the efficacy of a 3.2% glutaraldehyde product for disinfection of fibreoptic endoscopes with an automatic machine. J Hosp Infect 35:47–57. 270. Fraser VJ, Zuckerman G, Clouse RE, O’Rourke S, Jones M, Klasner J, Murray P. 1993. A prospective randomized trial comparing manual and automated endoscope disinfection methods. Infect Control Hosp Epidemiol 14:383– 389. 271. Centers for Disease Control and Prevention. 1999. Bronchoscopy-related infections and pseudoinfections—New York, 1996 and 1998. MMWR Morb Mortal Wkly Rep 48:557–560. 272. Bradley CR, Babb JR, Ayliffe GA. 1995. Evaluation of the Steris System 1 peracetic acid endoscope processor. J Hosp Infect 29:143–151. 273. Axon A, Jung M, Kruse A, Ponchon T, Rey JF, Beilenhoff U, Duforest-Rey D, Neumann C, Pietsch M, Roth K, Papoz A, Wilson D, Kircher-Felgenstreff I, Stief M, Blum R, Spencer KB, Mills J, Mart EP, Slowey B, Biering H, Lorenz U, et al. 2000. The European Society of Gastrointestinal Endoscopy (ESGE): check list for the purchase of washer-disinfectors for flexible endoscopes. Endoscopy 32:914–919. 274. Beilenhoff U, Neumann CS, Rey JF, Biering H, Blum R, Cimbro M, Kampf B, Rogers M, Schmidt V. 2008. ESGE-ESGENA guideline: cleaning and disinfection in gastrointestinal endoscopy. Endoscopy 40:939–957. 275. Farina A, Fievet MH, Plassart F, Menet MC, Thuillier A. 1999. Residual glutaraldehyde levels in fiberoptic endoscopes: measurement and implications for patient toxicity. J Hosp Infect 43:293–297. 276. Gurevich I, Dubin R, Cunha BA. 1996. Dental instrument and device sterilization and disinfection practices. J Hosp Infect 32:295–304. 277. Bagg J, Sweeney CP, Roy KM, Sharp T, Smith A. 2001. Cross infection control measures and the treatment of patients at risk of Creutzfeldt Jakob disease in UK general dental practice. Br Dent J 191:87–90. 278. Healy CM, Kearns HP, Coulter WA, Stevenson M, Burke FJ. 2004. Autoclave use in dental practice in the Republic of Ireland. Int Dent J 54:182–186. 279. Monarca S, Grottolo M, Renzi D, Paganelli C, Sapelli P, Zerbini I, Nardi G. 2000. Evaluation of environmental bacterial contamination and procedures to control cross infection in a sample of Italian dental surgeries. Occup Environ Med 57:721–726. 280. Piazza M, Borgia G, Picciotto L, Nappa S, Cicciarello S, Orlando R. 1995. Detection of hepatitis C virus-RNA by polymerase chain reaction in dental surgeries. J Med Virol 45:40–42. 281. ADA Council on Scientific Affairs and ADA Council on Dental Practice. 1996. Infection control recommendations for the dental office and the dental laboratory. J Am Dent Assoc 127:672–680. 282. Centers for Disease Control and Prevention. 1993. Recommended infection-control practices for dentistry. MMWR Morb Mortal Wkly Rep 42:1–12.

13. Decontamination, Disinfection, and Sterilization n 283. Rutala WA. 1996. Disinfection and sterilization of patient-care items. Infect Control Hosp Epidemiol 17:377– 384. 284. Scherrer M, Kümmerer K. 1997. Manual and automated processing of medical instruments. Environmental and economic aspects. Central Service 5:183–194. 285. Gurevich I, Tafuro P, Ristuccia P, Herrmann J, Young AR, Cunha BA. 1983. Disinfection of respirator tubing: a comparison of chemical versus hot water machine-assisted processing. J Hosp Infect 4:199–208. 286. Bradley CR, Fraise AP. 1996. Heat and chemical resistance of enterococci. J Hosp Infect 34:191–196. 287. U.S. Food and Drug Administation. 2002. Medical devices: classification for medical washer and medical washer disinfector. Fed Regist 67:69119–69121. 288. Borchardt MA, Bertz PD, Spencer SK, Battigelli DA. 2003. Incidence of enteric viruses in groundwater from household wells in Wisconsin. Appl Environ Microbiol 69:1172–1180. 289. Zuhlsdorf B, Kampf G, Floss H, Martiny H. 2005. Suitability of the German test method for cleaning efficacy in washer-disinfectors for flexible endoscopes according to prEN ISO 15883. J Hosp Infect 61:46–52. 290. Zuhlsdorf B, Martiny H. 2005. Intralaboratory reproducibility of the German test method of prEN ISO 15883– 1 for determination of the cleaning efficacy of washerdisinfectors for flexible endoscopes. J Hosp Infect 59:286– 291. 291. Joslyn LJ. 1991. Sterilization by heat, p 495–526. In Block SS (ed), Disinfection, Sterilization and Preservation. Lea & Febiger, Philadelphia, PA. 292. Graham GS. 1997. Decontamination: scientific principles, p 1–9. In Reichert M, Young JH (ed), Sterilization Technology. Aspen Publishers, Gaithersburg, MD. 293. Rutala WA, Gergen MF, Weber DJ. 1993. Evaluation of a rapid readout biological indicator for flash sterilization with three biological indicators and three chemical indicators. Infect Control Hosp Epidemiol 14:390–394. 294. Rutala WA, Jones SM, Weber DJ. 1996. Comparison of a rapid readout biological indicator for steam sterilization with four conventional biological indicators and five chemical indicators. Infect Control Hosp Epidemiol 17:423–428. 295. Bowie JH, Kennedy MH, Robertson I. 1975. Improved Bowie and Dick test. Lancet i:1135. 296. Vesley D, Nellis MA, Allwood PB. 1995. Evaluation of a rapid readout biological indicator for 121 degrees C gravity and 132 degrees C vacuum-assisted steam sterilization cycles. Infect Control Hosp Epidemiol 16:281–286. 297. Bryce EA, Roberts FJ, Clements B, MacLean S. 1997. When the biological indicator is positive: investigating autoclave failures. Infect Control Hosp Epidemiol 18:654– 656. 298. van Doornmalen JP, Dankert J. 2005. A validation survey of 197 hospital steam sterilizers in The Netherlands in 2001 and 2002. J Hosp Infect 59:126–130. 299. Widmer AF, Houston A, Bollinger E, Wenzel RP. 1992. A new standard for sterility testing for autoclaved surgical trays. J Hosp Infect 21:253–260. 300. Crow S. 1993. Steam sterilizers: an evolution in design. Infect Control Hosp Epidemiol 14:488–490. 301. Maki DG, Hassemer CA. 1987. Flash sterilization: carefully measured haste. Infect Control 8:307–310. 302. Rutala WA, Weber DJ, Chappell KJ. 1999. Patient injury from flash-sterilized instruments. Infect Control Hosp Epidemiol 20:458. 303. Favero MS, Manian FA. 1993. Is eliminating flash sterilization practical? Infect Control Hosp Epidemiol 14:479– 480. 304. Rutala WA, Gergen MF, Weber DJ. 1999. Sporicidal activity of a new low-temperature sterilization technology:

305. 306. 307.

308.

309.

310.

311.

312.

313.

314.

315. 316.

317.

318.

319.

320.

321.

322.

323.

324.

325.

215

the Sterrad 50 sterilizer. Infect Control Hosp Epidemiol 20:514–516. Daschner F. 1994. Steris System 1 in Germany. Infect Control Hosp Epidemiol 15:294, 296. Holton J, Shetty N. 1997. In-use stability of Nu-Cidex. J Hosp Infect 35:245–248. Hussaini SN, Ruby KR. 1976. Sporicidal activity of peracetic acid against B. anthracis spores. Vet Rec 98:257– 259. Wallace J, Agee PM, Demicco DM. 1995. Liquid chemical sterilization using peracetic acid. An alternative approach to endoscope processing. ASAIO J 41:151–154. Mannion PT. 1995. The use of peracetic acid for the reprocessing of flexible endoscopes and rigid cystoscopes and laparoscopes. J Hosp Infect 29:313–315. Kralovic RC. 1993. Use of biological indicators designed for steam or ethylene oxide to monitor a liquid chemical sterilization process. Infect Control Hosp Epidemiol 14:313–319. Gurevich I, Qadri SM, Cunha BA. 1993. False-positive results of spore tests from improper clip use with the STERIS chemical sterilant system. Am J Infect Control 21:42–43. Fuselier HAJ, Mason C. 1997. Liquid sterilization versus high level disinfection in the urologic office. Urology 50:337–340. Bruce ME, Will RG, Ironside JW, McConnell I, Drummond D, Suttie A, McCardle L, Chree A, Hope J, Birkett C, Cousens S, Fraser H, Bostock CJ. 1997. Transmissions to mice indicate that ‘new variant’ CJD is caused by the BSE agent. Nature 389:498–501. Kawashima T, Furukawa H, Doh-ura K, Iwaki T. 1997. Diagnosis of new variant Creutzfeldt-Jakob disease by tonsil biopsy. Lancet 350:68–69. Prusiner SB. 1997. Prion diseases and the BSE crisis. Science 278:245–251. Glatzel M, Stoeck K, Seeger H, Luhrs T, Aguzzi A. 2005. Human prion diseases: molecular and clinical aspects. Arch Neurol 62:545–552. Hill AF, Desbruslais M, Joiner S, Sidle KC, Gowland I, Collinge J, Doey LJ, Lantos P. 1997. The same prion strain causes vCJD and BSE. Nature 389:448–450. Aguzzi A, Weissmann C. 1998. Spongiform encephalopathies. The prion’s perplexing persistence. Nature 392:763–764. Blattler T, Brandner S, Raeber AJ, Klein MA, Voigtlander T, Weissmann C, Aguzzi A. 1997. PrP-expressing tissue required for transfer of scrapie infectivity from spleen to brain. Nature 389:69–73. Sailer A, Bueler H, Fischer M, Aguzzi A, Weissmann C. 1994. No propagation of prions in mice devoid of PrP. Cell 77:967–968. Chazot G, Broussolle E, Lapras C, Blattler T, Aguzzi A, Kopp N. 1996. New variant of Creutzfeldt-Jakob disease in a 26-year-old French man. Lancet 347:1181. Valleron AJ, Boelle PY, Will R, Cesbron JY. 2001. Estimation of epidemic size and incubation time based on age characteristics of vCJD in the United Kingdom. Science 294:1726–1728. Gill ON, Spencer Y. 2013. Prevalent abnormal prion protein in human appendixes after bovine spongiform encephalopathy epizootic: large scale survey. BMJ 347:f5675. Herzog C, Sales N, Etchegaray N, Charbonnier A, Freire S, Dormont D, Deslys JP, Lasmezas CI. 2004. Tissue distribution of bovine spongiform encephalopathy agent in primates after intravenous or oral infection. Lancet 363:422–428. Klein MA, Frigg R, Flechsig E, Raeber AJ, Kalinke U, Bluethman H, Bootz F, Suter J, Zinkernagel RM, Aguzzi

216 n

326.

327.

328.

329. 330. 331.

332. 333.

334. 335.

336.

337.

338.

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

A. 1997. A crucial role for B cells in neuroinvasive scrapie. Nature 390:687. Heikenwalder M, Zeller N, Seeger H, Prinz M, Klohn PC, Schwarz P, Ruddle NH, Weissmann C, Aguzzi A. 2005. Chronic lymphocytic inflammation specifies the organ tropism of prions. Science 307:1107–1110. Aguzzi A, Glatzel M. 2004. vCJD tissue distribution and transmission by transfusion—a worst-case scenario coming true? Lancet 363:411–412. Kampf G, Bloss R, Martiny H. 2004. Surface fixation of dried blood by glutaraldehyde and peracetic acid. J Hosp Infect 57:139–143. Taylor DM. 1999. Inactivation of prions by physical and chemical means. J Hosp Infect 43(Suppl):S69–S76. Dormont D. 1996. How to limit the spread of CreutzfeldtJakob disease. Infect Control Hosp Epidemiol 17:521–528. Taylor DM, Woodgate SL, Fleetwood AJ, Cawthorne RJ. 1997. Effect of rendering procedures on the scrapie agent. Vet Rec 141:643–649. Charatan F. 2001. United States takes precautions against BSE. West J Med 174:235. World Health Organization. 1983. Laboratory Safety Manual. World Health Organization, Geneva, Switzerland. Larson E. 1988. Guideline for use of topical antimicrobial agents. Am J Infect Control 16:253–266. DiazGranados CA, Jones MY, Kongphet-Tran T, White N, Shapiro M, Wang YF, Ray SM, Blumberg HM. 2009. Outbreak of Pseudomonas aeruginosa infection associated with contamination of a flexible bronchoscope. Infect Control Hosp Epidemiol 30:550–555. Shimono N, Takuma T, Tsuchimochi N, Shiose A, Murata M, Kanamoto Y, Uchida Y, Morita S, Matsumoto H, Hayashi J. 2008. An outbreak of Pseudomonas aeruginosa infections following thoracic surgeries occurring via the contamination of bronchoscopes and an automatic endoscope reprocessor. J Infect Chemother 14:418– 423. Bou R, Aguilar A, Perpinan J, Ramos P, Peris M, Lorente L, Zuniga A. 2006. Nosocomial outbreak of Pseudomonas aeruginosa infections related to a flexible bronchoscope. J Hosp Infect 64:129–135. Cetre JC, Nicolle MC, Salord H, Perol M, Tigaud S, David G, Bourjault M, Vanhems P. 2005. Outbreaks of contaminated broncho-alveolar lavage related to intrinsically defective bronchoscopes. J Hosp Infect 61:39–45.

339. Fraser TG, Reiner S, Malczynski M, Yarnold PR, Warren J, Noskin GA. 2004. Multidrug-resistant Pseudomonas aeruginosa cholangitis after endoscopic retrograde cholangiopancreatography: failure of routine endoscope cultures to prevent an outbreak. Infect Control Hosp Epidemiol 25:856–859. 340. Sorin M, Segal-Maurer S, Mariano N, Urban C, Combest A, Rahal JJ. 2001. Nosocomial transmission of imipenem-resistant Pseudomonas aeruginosa following bronchoscopy associated with improper connection to the Steris System 1 processor. Infect Control Hosp Epidemiol 22:409–413. 341. Rutala WA, Weber DJ. 1999. Disinfection of endoscopes: review of new chemical sterilants used for high-level disinfection. Infect Control Hosp Epidemiol 20:69–76. 342. Bronowicki JP, Venard V, Botte C, Monhoven N, Gastin I, Chone L, Hudziak H, Rhin B, Delanoe C, LeFaou A, Bigard MA, Gaucher P. 1997. Patient-to-patient transmission of hepatitis C virus during colonoscopy. N Engl J Med 337:237–240. 343. Michele TM, Cronin WA, Graham NM, Dwyer DM, Pope DS, Harrington S, Chaisson RE, Bishai WR. 1997. Transmission of Mycobacterium tuberculosis by a fiberoptic bronchoscope. Identification by DNA fingerprinting. JAMA 278:1093–1095. 344. Agerton T, Valway S, Gore B, Pozsik C, Plikaytis B, Woodley C, Onorato I. 1997. Transmission of a highly drug-resistant strain (strain W1) of Mycobacterium tuberculosis. Community outbreak and nosocomial transmission via a contaminated bronchoscope. JAMA 278:1073– 1077. 345. Blanc DS, Parret T, Janin B, Raselli P, Francioli P. 1997. Nosocomial infections and pseudoinfections from contaminated bronchoscopes: two-year follow up using molecular markers. Infect Control Hosp Epidemiol 18:134– 136. 346. Gubler JG, Salfinger M, von Graevenitz A. 1992. Pseudoepidemic of nontuberculous mycobacteria due to a contaminated bronchoscope cleaning machine. Report of an outbreak and review of the literature. Chest 101:1245–1249. 347. Carbonne A, Thiolet JM, Fournier S, Fortineau N, Kassis-Chikhani N, Boytchev I, Aggoune M, Seguier JC, Senechal H, Tavolacci MP, Coignard B, Astagneau P, Jarlier V. 2010. Control of a multi-hospital outbreak of KPC-producing Klebsiella pneumoniae type 2 in France, September to October 2009. Euro Surveill 15:19734.

Biothreat Agents SUSAN E. SHARP AND MICHAEL LOEFFELHOLZ

14 Through a collaborative effort involving the Federal Bureau of Investigation and the Association of Public Health Laboratories (APHL), the LRN was established in August 1999. Its objective was to improve and ensure an effective laboratory response to bioterrorism. The LRN worked toward this goal by improving the public health infrastructure, increasing laboratory capacity and staffing, and training laboratory personnel in advanced diagnostic technologies. The LRN forms an integrated network of more than 150 local and state public health, federal, and military laboratories (representing all 50 states, Australia, Canada, the United Kingdom, Mexico, and South Korea) that can respond to bioterrorism, chemical terrorism, and other public health emergencies. The LRN states its mission as follows: “The LRN and its partners will develop, maintain and strengthen an integrated national and international network of laboratories that can respond quickly to needs for rapid testing, timely notification and secure messaging of results associated with acts of biological or chemical terrorism and other high priority public health emergencies” (http:// www.bt.cdc.gov/lrn/). There are an estimated 25,000 private and commercial laboratories (including clinical, food testing, veterinary diagnostic, and environmental testing laboratories) in the United States, some of which serve as critical sentinel laboratories. While most of these laboratories do not perform confirmatory testing, they represent the first contact with patients and are in a position to alert public health officials of possible biothreat agents. These laboratories conduct tests to rule out other diseases and send possible biothreat samples/organisms to the appropriate LRN reference laboratories for confirmatory testing. The LRN structure is shown in Fig. 1. LRN sentinel laboratories are certified to perform highcomplexity testing under the Clinical Laboratory Improvement Amendments of 1988 (CLIA) by the Centers for Medicare and Medicaid Services (CMS) for the applicable microbiology specialty. Laboratory in-house testing includes Gram stains and at least one of the following: lower respiratory tract, wound, or blood culture. LRN reference laboratories are responsible for investigation and/or referral of specimens to LRN national laboratories. LRN reference laboratories consist of over 100 state and local public health, military, federal, and international laboratories (veterinary, agriculture, food, and water testing laboratories) in and outside the United States. LRN national laboratories, including

The ideal qualities for a successful biothreat agent are a high rate of illness in exposed persons/animals, a high case fatality rate, a short incubation period, and a paucity of immunity in the targeted population. In addition, success is also influenced by the availability of treatment, the ability of the agent to transmit from person to person, the ease with which the agent can be produced and disseminated, and a disease that is difficult, at least initially, to recognize clinically or to diagnose. General clues that one may be dealing with an unrecognized biothreat event include a large outbreak of illness with a high death rate, a recognized case(s) of an uncommon disease, illness in a geographic region where the disease is not endemic, disease out of its usual seasonality, simultaneous outbreaks of the same disease in various parts of the country/ world, and the presence of sick and dying animals. The Centers for Disease Control and Prevention (CDC) have classified potential biothreat agents into categories depending on the basis of their threat to national security, with those organisms belonging to category A being the most serious threats, due to their unique features in causing diseases capable of mass destruction (Table 1). These and other agents are discussed in this chapter.

The LRN The Laboratory Response Network (LRN) was established through the work of the Department of Health and Human Services (HHS) and the CDC, in accordance with Presidential Decision Directive 39. This directive, signed by President Bill Clinton on 22 May 1998, made the fight against terrorism a top national security priority and defined policies regarding the federal response to threats or acts of terrorism involving nuclear, biological, and/or chemical material and/ or weapons of mass destruction. Specifically, this directive outlined the United States’ efforts to “deter, defeat and respond vigorously to all terrorist attacks on our territory and against our citizens, or facilities, whether they occur domestically, in international waters or airspace or on foreign territory. The United States regards all such terrorism as a potential threat to national security as well as a criminal act and will apply all appropriate means to combat it. In doing so, the U.S. shall pursue vigorously efforts to deter and preempt, apprehend and prosecute, or assist other governments to prosecute, individuals who perpetrate or plan to perpetrate such attacks” (http://www.fas.org/irp/off docs/pdd39.htm).

doi:10.1128/9781555817381.ch14

217

218 n TABLE 1

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS HHS and USDA select agents and toxins (with biothreat categories)a,b

HHS select agents and toxins Abrin Botulinum neurotoxins* (A) Botulinum neurotoxin-producing species of Clostridium* (A) Conotoxins (short, paralytic alpha conotoxins containing the following amino acid sequence: X1CCX2PACGX3X4X5X6CX7) Coxiella burnetii (B) Crimean-Congo hemorrhagic fever virus Diacetoxyscirpenol Eastern equine encephalitis virus (B) Ebola virus* (A) Francisella tularensis* (A) Lassa fever virus (A) Lujo virus Marburg virus* (A) Monkeypox virus Reconstructed replication-competent forms of the 1918 pandemic influenza virus containing any portion of the coding regions of all eight gene segments (reconstructed 1918 influenza virus) Ricin (B) Rickettsia prowazekii Severe acute respiratory syndrome-associated coronavirus (SARS-CoV) Saxitoxin South American hemorrhagic fever viruses (Chapare, Guanarito, Junin, Machupo, and Sabia viruses) Staphylococcal enterotoxin A, B, C, D, and E subtypes (B) T-2 toxin Tetrodotoxin Tick-borne encephalitis complex (flavi-) viruses (Far Eastern subtype, Siberian subtype) (C) Kyasanur Forest disease virus Omsk hemorrhagic fever virus Variola major virus (smallpox virus)* (A) Variola minor virus (alastrim)* Yersinia pestis* (A)

Overlapping select agents and toxins Bacillus anthracis* (A) Bacillus anthracis Pasteur strain Brucella abortus (B) Brucella melitensis (B) Brucella suis (B) Burkholderia mallei* (B) Burkholderia pseudomallei* Hendra virus Nipah virus (C) Rift Valley fever virus Venezuelan equine encephalitis virus

Other critical agents for public health preparedness Western equine encephalitis virus (B) Epsilon toxin of Clostridium perfringens (B) Salmonella species (B) Shigella dysenteriae (B) Escherichia coli O157:H7 (B) Vibrio cholerae (B) Cryptosporidium parvum (B) Hantavirus (C) Yellow fever virus (C) Multidrug-resistant Mycobacterium tuberculosis (C)

a *, Tier 1 agent. Select agent regulations were revised in October 2012 to designate “tier 1” agents with a documented risk of causing a high-consequence event higher than those with other biological select agents and toxins. Criteria for tier 1 status were as follows: (i) ability to produce a mass casualty event or devastating effects to the economy, (ii) communicability, (iii) low infectious dose, and (iv) history of or current interest in weaponization based on threat reporting. See http:// www.selectagents.gov/select%20agents%20and%20toxins%20list.html. b Note that the CDC divides bioterrorism agents into three categories (A, B, and C) depending on how easily they can be spread and the severity of illness or rate of death they cause. Category A agents are considered the highest risk, as they can easily be disseminated or transmitted from person to person, result in high mortality rates, have the potential for major public health impacts, might cause public panic and social disruption, and require special action for public health preparedness. Category B agents are the second-highest-priority agents and include those that are moderately easy to disseminate, result in moderate morbidity rates and low mortality rates, and require specific enhancements of the CDC’s diagnostic capacity and enhanced disease surveillance. Category C agents are the thirdhighest-priority agents and include emerging pathogens that could be engineered for mass dissemination in the future because of availability, ease of production and dissemination, and potential for high morbidity and mortality rates and major health impacts.

those operated by the CDC and the U.S. Army Medical Research Institute of Infectious Diseases (USAMRIID), have specialized infrastructure, training, and biosecurity that are necessary for handling highly infectious biological agents. These laboratories are responsible for specialized strain characterizations, susceptibility testing, and bioforensics. The American Society for Microbiology (ASM), along with the APHL and the CDC, has been instrumental in the development of sentinel-level laboratory testing

protocols for the detection of biothreat agents. These protocols were developed to allow LRN sentinel laboratories to rapidly rule out or refer suspected biothreat agents to the appropriate LRN reference laboratory. Along with the CDC and APHL, ASM serves as a resource for training and education for microbiologists in hospital and commercial laboratories regarding their roles and responsibilities as LRN sentinel laboratories. These protocols and other useful information regarding bioterror agents

14. Biothreat Agents n

219

The USDA’s Animal and Plant Health Inspection Service (APHIS) then established the Agricultural Select Agent Program. These two programs constitute the Federal Select Agent Program. The Program greatly enhances the nation’s oversight of the safety and security of select agents by developing, implementing, and enforcing the Select Agent Regulations, maintaining a national database, inspecting entities that possess, use, or transfer select agents, and ensuring that all individuals who work with these agents undergo a security risk assessment performed by the Federal Bureau of Investigation/Criminal Justice Information Service. The list of agents that the Federal Select Agent Program regulates is shown in Table 1. The list is reviewed by the CDC and APHIS at least every 2 years to determine if agents need to be added to or deleted from the list. Most recently (2012), Coccidioides species were deleted from the select agent list. Refer to the ASM website for flowcharts and additional information on bioterror agents and preparedness: http://www.asm.org/index.php/issues/ sentinel-laboratory-guidelines. FIGURE 1 Structure of the Laboratory Response Network. doi:10.1128/9781555817381.ch14.f1

can be found at the following ASM website page: http:// www.asm.org/index.php/guidelines/sentinel-guidelines. The College of American Pathologists (CAP), together with the CDC, also serves an important role in promoting the diagnostic capability and capacity of clinical and public health laboratories by developing the Laboratory Preparedness Exercise (LPX) proficiency test. There are additional sources of proficiency testing for laboratory bioterrorism preparedness, such as the Wisconsin State Laboratory of Hygiene (http://www.slh.wisc.edu/profi ciency/). These proficiency programs help to ensure the capability of participating sentinel laboratories to rule out and/or refer biological threat agents to the appropriate LRN reference laboratory. A directory of LRN public health reference laboratories is available at http://www.cdc.gov/mmwr/international/ relres.html.

Federal Select Agent Program The Federal Select Agent Program was established due to a heightened concern about the ease with which diseasecausing agents could be obtained. Section 511 of the Antiterrorism and Effective Death Penalty Act of 1996 (Public Law 104-132) directed the HHS to establish a list of biological agents and toxins that could threaten public health and safety, procedures for governing the transfer of those agents, and training requirements for entities working with these “select agents.” HHS delegated the authority to implement this Act to the CDC, which then established the CDC Select Agent Program. Congress significantly strengthened oversight of select agents after the anthrax attacks of 2001 by passing the USA Patriot Act and the Bioterrorism Act, which together restricted access to select agents and increased safeguards and security measures, as well as oversight of the possession and use of select agents. In addition to strengthening the regulatory authorities of the CDC, the Bioterrorism Act also granted comparable regulatory authorities to the U.S. Department of Agriculture (USDA) over select agents that pose a severe threat to animal and plant health or products.

CDC Bioterrorism Agents and Diseases: Categories and Definitions The CDC divides bioterrorism agents into three categories (A, B, and C) depending on how easily they can be spread and the severity of illness or rate of death they cause (Table 1) (http://www.bt.cdc.gov/bioterrorism/over view.asp). Category A agents are considered the highest risk, as they can easily be disseminated or transmitted from person to person, result in high mortality rates, have the potential for major public health impacts, might cause public panic and social disruption, and require special action for public health preparedness. Agents included in this category are Bacillus anthracis, Clostridium botulinum toxin, Yersinia pestis, variola virus (the agent of smallpox), Francisella tularensis, and hemorrhagic fever viruses (HFVs), such as filoviruses (e.g., Ebola virus and Marburg virus) and arenaviruses (e.g., Lassa virus and Machupo virus). Characteristics of several of these agents are listed in Table 2. Category B agents are the second-highest-priority agents and include those that are moderately easy to disseminate, result in moderate morbidity rates and low mortality rates, and require specific enhancements of the CDC’s diagnostic capacity and enhanced disease surveillance. Category B agents include Brucella species (Brucella abortus, Brucella melitensis, and Brucella suis), epsilon toxin of Clostridium perfringens, food-safety threats (e.g., Salmonella species, Escherichia coli O157:H7, and Shigella species), Burkholderia mallei, Burkholderia pseudomallei, Chlamydia psittaci, Coxiella burnetii, ricin toxin from Ricinus communis (castor beans), staphylococcal enterotoxin B (SEB), Rickettsia prowazekii, viral encephalitis viruses (such as alphaviruses, e.g., Venezuelan equine encephalitis [VEE] virus, Eastern equine encephalitis [EEE] virus, and Western equine encephalitis [WEE] virus), and water-safety threats (e.g., Vibrio cholerae and Cryptosporidium parvum). Category C agents are the third-highest-priority agents and include emerging pathogens that could be engineered for mass dissemination in the future because of availability, ease of production and dissemination, and potential for high morbidity and mortality rates and major health impacts. These emerging pathogens are covered in their individual organism chapters.

220 TABLE 2

Summary of biothreat agent characteristics Burkholderia pseudomallei and B. mallei

B. anthracis

Y. pestis

Large Gram-positive rod, nonmotile. From blood agar (BA): no capsule, central to subterminal spores that do not enlarge the cell. From blood: capsule, no spores.

Plump Gram-negative rod. Gram stain: ± bipolar or “safety pin” appearance. Wright-Giemsa stain: bipolar or “safety pin” appearance.

Growth

Standard conditions, extremely rapid.

28°C optimum, without agitation; 35–37°C, more slowly.

Colonial morphology (BA)

Nonhemolytic, ground glass, irregular/wavy edges, tenacious, “beaten egg whites” when touched with loop.

Nonhemolytic, pinpoint colonies at 24–48 h, “fried egg” or “hammered copper” or shiny at 48–72 h.

B. pseudomallei: small, smooth creamy colonies in first 1 to 2 days, gradually changing after a few days to dry, wrinkled colonies similar to those of Pseudomonas stutzeri. B. mallei: smooth, gray, translucent colonies without pigment.

Tests

Catalase (+) Catalase (+), oxidase (−), urease (−). On MacConkey agar (MAC): lactose (−), indole (−).

Catalase (+), colistin (10 μg) and polymyxin B (300 U) (resistant), motility (+, B. pseudomallei; −, B. mallei), indole (−), oxidase (+, B. pseudomallei; ±, B. mallei). On MAC: lactase (−, B. pseudomallei; − or no growth, B. mallei).

Characteristic Gram stain morphology

B. pseudomallei: small Gramnegative rod. B. mallei: small Gramnegative coccobacillus. Gram stain: ± bipolar or “safety pin” appearance (B. pseudomallei). Wright-Giemsa stain: bipolar or “safety pin” appearance (B. pseudomallei). 35–37°C, ambient atmosphere, though CO2 is acceptable.

F. tularensis

Brucella spp.

Minute Gram-negative coccobacillus, poorly staining, smaller than Haemophilus influenzae, pleomorphic.

Tiny Gram-negative coccobacillus, faintly staining.

Aerobic conditions. Growth is best on media containing cysteine, such as BCYE, but will often grow initially on chocolate agar or BA. Does not passage well on BA.

Grows in blood culture media, can require blind subculturing.

Catalase (weakly +), oxidase (−), β-lactamase (+), satellites (−). On MAC: no growth.

Oxidase (+), catalase (+), urease (+) (though some strains are negative), satellites (−). On MAC: poor to no growth.

Variola virus (smallpox)

Grows in most cell lines, unusual or unrecognizable cytopathic effect (CPE)

Small colonies, punctate after 48 h, nonhemolytic.

CPE can be passed.

14. Biothreat Agents n 221

BIOTHREAT AGENTS AND INFECTIONS Category A Agents Anthrax (Bacillus anthracis) Significance The biothreat associated with the inhalation of spores from B. anthracis has been known for decades, and in the past the organism was developed as a potential military weapon by many countries, including the United States (1, 2). In 2001, envelopes containing confirmed anthrax spores were mailed to southern Florida, New York City, and Washington, D.C., resulting in 22 cases of anthrax (11 inhalational cases, with 5 deaths, and 11 cases of cutaneous anthrax) (2).

Plague (Yersinia pestis) Significance The first reported use of Y. pestis as a bioweapon was in the 1300s, as the Tartars, invading the city of Kaffa (now Feodosiya, Ukraine), catapulted bodies of plague victims over the city walls, resulting in an epidemic (4). Many countries have developed this agent as a bioweapon, including the former Soviet Union and Japan (5). An incident involving a white supremacist who fraudulently obtained cultures of Y. pestis from the American Type Culture Collection prompted the U.S. Congress to pass a 1996 regulation governing the acquisition, transfer, and use of agents that could be used for bioterrorism purposes (6).

Transmission and Disease Transmission and Disease B. anthracis is a large, aerobic or facultative anaerobic, spore-forming, Gram-positive bacillus that naturally inhabits the soil. Four forms of the disease can occur: cutaneous (>95% of all human anthrax cases), inhalational, gastrointestinal, and meningitis forms. The inhalational form (acquired through inhalation of organism spores) is primarily associated with bioterror attacks and causes pulmonary anthrax. In addition, anthrax meningitis can occur as a result of a bioterror event. Any case of inhalational or meningeal anthrax should prompt suspicion of possible bioterrorism. Human-to-human transmission of anthrax has not been reported. See ASM’s sentinel-level clinical microbiology laboratory guidelines for anthrax (http://www.asm.org/ index.php/guidelines/sentinel-guidelines). Additional information regarding B. anthracis can be found in chapter 26.

Botulism (Clostridium botulinum) Significance Clostridial toxins are among the most powerful neurotoxins known. Botulinum toxin acts at the neuromuscular junction and at the peripheral autonomic synapses, resulting in neuromuscular weakness and autonomic dysfunction (3). A bioterror attack using botulinum toxin could occur through ingestion or inhalation of the toxin, such as contamination of a food or air supply. Many countries, including Japan, the United States, the former Soviet Union, and Iraq, have, at some point, developed botulinum toxin for biological warfare (3).

Transmission and Disease C. botulinum is an anaerobic, spore-forming, Gram-positive bacillus that is found naturally in the environment (soil and water). The organisms primarily associated with human disease produce toxin types A, B, and E. Botulism is naturally acquired from contaminated food products processed by methods that do not destroy spores. This allows bacteria contained in the food to become vegetative and toxins to be produced. The clinical presentation associated with a bioterror attack with botulinum would include gastrointestinal symptomology (nausea, vomiting, and diarrhea) followed by neurological signs of dry mouth and blurred and/ or double vision. In cases of severe disease, symptoms include difficulty swallowing, voice impairment, and peripheral muscle weakness. If the muscles of the respiratory tract are involved, respiratory failure and death may result. See ASM’s sentinel-level clinical microbiology laboratory guidelines for botulism (http://www.asm.org/index.php/ guidelines/sentinel-guidelines). Additional information regarding C. botulinum can be found in chapter 53.

Y. pestis is a slow-growing (optimal growth is at 28°C; up to 5 days of incubation is recommended), plump, Gramnegative coccobacillus/bacillus and is part of the Enterobacteriaceae family. Plague is a worldwide zoonotic disease that is transmitted between animals via infected fleas. There are three forms of plague: bubonic, septicemic, and pneumonic. The pneumonic form would be the most likely disease manifestation seen in the event of a bioterror attack. See ASM’s sentinel-level clinical microbiology laboratory guidelines for plague (http://www.asm.org/index.php/guidelines/sentinelguidelines). Additional information regarding Y. pestis can be found in chapter 39.

Smallpox (Variola Major) Significance An excellent overview of smallpox and its historical use as a biothreat weapon is given by Rotz et al. (7). Briefly, one of the first possibilities of smallpox being used as a biothreat weapon dates back to the 1600s, when the Spanish supplied the natives of South America with smallpoxcontaminated clothing in an effort to gain control of their lands. Then, in the 1760s, the North American British forces gave smallpox-contaminated blankets to the native Indians in an effort to reduce their populations and gain control of land. In more recent times, the Russians associated with Bioperparat worked with many potential biothreat agents, including smallpox, for use in intercontinental missiles (8). Today, concern still exists regarding illegitimate stores of the agent and their potential use in bioterrorism, especially because routine vaccination has not been in place for decades, leading to a susceptible populace.

Transmission and Disease Variola virus belongs to the genus Orthopoxvirus in the family Poxviridae. Variola virus is a large, brick-shaped virus measuring approximately 302 to 350 nm by 244 to 270 nm and is comprised of a single linear double-stranded DNA genome. As there is no known disease associated with smallpox in the world, even one case would prompt the suspicion of a bioterror incident. Human transmission occurs primarily by inhalation of airborne particles containing the virus, but transmission may occur via fomites (contaminated bedding or clothing) as well. Disease consists of symptoms such as malaise, fever, chills, vomiting, headache, and backache, with the eventual formation of a papular rash on the face, hands, and arms, later spreading to the legs and, lastly, the trunk. The mortality due to smallpox in the unvaccinated population is approximately 30% (9). See ASM’s sentinel-level clinical microbiology laboratory guidelines for unknown viruses (http://

222 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

www.asm.org/index.php/guidelines/sentinel-guidelines). Additional information regarding smallpox can be found in chapter 107.

Tularemia (Francisella tularensis) Significance The highly infectious nature of F. tularensis, due to low inhalation inocula and a very susceptible human population, as well as its substantial morbidity and mortality in untreated patients, makes this bacterium a potential agent of bioterrorism. The Japanese began experiments on human prisoners for use of this agent as a military weapon in the early 1930s, and in 1955 the United States used volunteers and military personnel to test the effects of inhaled organisms (10). In the 1990s, Russia developed antibiotic- and vaccineresistant strains of F. tularensis (http://www.pbs.org/wgbh/ nova/bioterror/agen_tularemia.html).

Transmission and Disease F. tularensis is an extremely small, fastidious, pleomorphic, Gram-negative coccobacillus. Aerosolized particles containing the organism account for the acquisition of pneumonic tularemia, the primary disease that would be associated with a bioterror attack; however, patients inhaling aerosolized organisms may present with other forms of tularemia (ulceroglandular, glandular, oculoglandular, pharyngeal, and typhoidal). Respiratory failure or shock causes most fatalities from pneumonic disease (http://www.pbs.org/ wgbh/nova/bioterror/agen_tularemia.html). Two subspecies of F. tularensis, F. tularensis subsp. tularensis (type A) and F. tularensis subsp. holarctica (type B), are of clinical importance in causing tularemia. See ASM’s sentinel-level clinical microbiology laboratory guidelines for F. tularensis (http://www.asm.org/index.php/guidelines/sentinelguidelines). Additional information regarding F. tularensis can be found in chapter 46.

Diseases Caused by HFVs Significance Hemorrhagic fever viruses (HFVs) were developed as biological weapons by both the United States and the former Soviet Union. The United States stopped this program and destroyed all weapons in the late 1960s.

Transmission and Disease The HFVs comprise four families: the Arenaviridae, Bunyaviridae, Filoviridae, and Flaviviridae. The clinical course following exposure varies with the virus causing the infection, but typically patients have symptoms of fever, myalgia, and malaise followed by exacerbation of symptoms and development of prostration and hemorrhage, with central nervous system (CNS) depression. Patients who develop shock with extensive hemorrhage and CNS damage generally have poor outcomes (11). See ASM’s sentinel-level clinical microbiology laboratory guidelines for unknown viruses (http://www.asm.org/ index.php/guidelines/sentinel-guidelines). Additional information regarding HFVs can be found in section IV of this Manual.

mid-1950s. Brucella species are considered among the agents less likely to be utilized in a bioterrorism attack, in part because brucellosis results in a high morbidity but low mortality. However, it remains a threat because the disease process is long and can be incapacitating. Brucella species listed as category B agents are B. abortus, B. melitensis, and B. suis.

Transmission and Disease Brucellosis is a zoonotic infection, with four species being recognized as causing infection in humans: B. abortus (cattle), B. melitensis (goats, sheep, and camels), B. suis (pigs), and Brucella canis (dogs). In addition, it is a common laboratory-acquired infection. Brucella organisms are small, aerobic, Gram-negative coccobacilli that grow slowly (2 to 3 days for initial isolation). Brucella can cause both acute and chronic infections. Brucellosis is a disease of nonspecific symptoms, consisting of fever, sweats, headache, anorexia, back pain, and weight loss, which can last for months and relapse after discontinuation of therapy (12). The chronic form of the disease can mimic miliary tuberculosis, with suppurative lesions in the liver, spleen, and bone. Brucellosis has a mortality of 5% in untreated individuals. See ASM’s sentinel-level clinical microbiology laboratory guidelines for Brucella (http://www.asm.org/index.php/guidelines/ sentinel-guidelines). Additional information regarding Brucella species can be found in chapter 47.

Glanders and Melioidosis (Burkholderia mallei and B. pseudomallei) Significance It is believed that B. mallei was used to infect large numbers of Russian horses and mules on the Eastern Front during World War I and that the Japanese infected horses, civilians, and prisoners of war during World War II. The United States studied this agent as a possible biothreat agent in 1943 and 1944 but did not weaponize it. The former Soviet Union is also believed to have been interested in B. mallei as a potential biothreat agent (13).

Transmission and Disease Glanders and melioidosis are related diseases produced by bacteria of the Burkholderia species, which are nonsporulating, obligately aerobic, Gram-negative bacilli. They produce similar diseases which consist of several forms: localized infections, pulmonary infections (pneumonia, pulmonary abscesses, and pleural effusions), septicemia, and a chronic form (multiple abscesses in internal organs). Both B. mallei and B. pseudomallei are considered potential biothreat agents in the aerosolized/pulmonary form in that they are highly infectious by inhalation and because of their resistance to many routine antibiotics (http:// emedicine.medscape.com/article/830235-overview). See ASM’s sentinel-level clinical microbiology laboratory guidelines for Burkholderia species (http://www.asm.org/ index.php/guidelines/sentinel-guidelines). Additional information regarding Burkholderia species can be found in chapter 43.

Q Fever (Coxiella burnetii) Category B Agents Brucellosis (Brucella Species) Significance B. suis was the first biological agent to be utilized in the biological warfare program in the United States during the

Significance C. burnetii has a very low infective dose and is resistant to the effects of drying and heat. Humans are very susceptible to infection with this organism, and a single organism is capable of causing disease in a susceptible person. In the

14. Biothreat Agents n 223

1960s, the U.S. military considered Coxiella an excellent “incapacitating” agent, as the disease is debilitating but rarely lethal, and envisioned using this agent to cripple enemy forces. Russia, and possibly also Iraq, has also developed and tested the Q fever agent as a biothreat agent, and the cult Aum Shinrikyo obtained C. burnetii but was unsuccessful in weaponizing it.

Transmission and Disease C. burnetii is a pleomorphic, Gram-negative, intracellular coccobacillus that cannot be cultured on routine bacteriologic media. Q fever is a zoonotic disease seen primarily in parturient goats, sheep, and cattle. Aerosolized contaminated dust particles are the source of human infection, and the infectious dose is very low. There are three clinical presentations of this infection: atypical pneumonia, progressive pneumonia (of a rapid nature), and as an incidental finding in a patient with fever and pneumonia. The last description is the most common form of the disease (14). Q fever may present as a chronic infection. See ASM’s sentinel-level clinical microbiology laboratory guidelines for Q fever (http://www.asm.org/index.php/guidelines/sentinelguidelines). Additional information regarding Q fever can be found in chapter 66.

Staphylococcal Enterotoxins Significance Staphylococcal enterotoxin B (SEB) was part of the United States’ bioweapons program until the early 1970s. SEB was thought to be useful as a weapon due to its ease of aerosolization, stability, and ability to produce multisystem organ failure and death if inhaled in large amounts. Even if inhaled in smaller amounts, this toxin produces a debilitating disease that takes up to 2 weeks for recovery.

where it causes edema and hemorrhage. It is fast acting, causing death within hours or minutes of exposure (17). Documented human cases of enterotoxemia due to epsilon toxin are very rare, with only two reported cases, both reported in 1955 (16). Because of its rarity, any case of enterotoxemia due to epsilon toxin should prompt suspicion of possible bioterrorism. As a biothreat agent, epsilon toxin would likely be delivered as an aerosol or through water- or foodborne routes. Additional information regarding C. perfringens and epsilon toxin can be found in chapter 53.

Psittacosis (Chlamydia psittaci) Significance C. psittaci survives for long periods in dry environments, is spread naturally through the aerosol route, and has the potential to cause severe disease in humans. For these reasons, it is listed as a potential bioterrorism agent.

Transmission and Disease C. psittaci is an obligately intracellular bacterium that causes respiratory disease in birds. Human infections (psittacosis) are usually associated with exposure to bird secretions. Gardening and lawn mowing have been linked with psittacosis, and person-to-person transmission may occur (18). C. psittaci infection in humans usually presents as a mild to severe respiratory infection, with a wide range of symptoms. Signs and symptoms include fever, headache, muscular aches, nonproductive cough, and difficulty breathing. Extrapulmonary complications may also occur. As a biothreat agent, C. psittaci would likely be spread through the aerosol route. Additional information regarding C. psittaci can be found in chapter 63.

Epidemic Typhus (Rickettsia prowazekii) Transmission and Disease Significance SEB is one of several enterotoxins produced by Staphylococcus aureus, which is a ubiquitous, nonmotile, Grampositive coccus found on the skin and mucous membranes of humans and animals. SEB is also a potent superantigen capable of massive overstimulation of the immune system, resulting in an overwhelming inflammatory response and an endotoxin-like shock which can result in multisystem organ failure and death (15). See ASM’s sentinel-level clinical microbiology laboratory guidelines for staphylococcal enterotoxin B (http://www.asm.org/index.php/guidelines/ sentinel-guidelines). Additional information regarding SEB can be found in chapter 21.

Epsilon Toxin (Clostridium perfringens) Significance Because of its potency—nearly equal to that of botulinum neurotoxin (16)—epsilon toxin of C. perfringens is considered a potential bioterrorism agent.

Among the characteristics that make R. prowazekii a potential bioterrorism agent are its aerosol transmission, high infectivity, stable extracellular forms, and high virulence. R. prowazekii was developed as a biological weapon by several countries during the 1930s and 1940s (19).

Transmission and Disease R. prowazekii is an obligately intracellular bacterium that relies on an arthropod host (human body louse) as its principal vector. The classic epidemiologic cycle of epidemic typhus requires the feeding of an infected louse on a susceptible human host. Symptoms include high fever, myalgia, and arthralgia. Because of the nonspecific symptoms, infections may go undiagnosed or be treated with ineffective antibiotics. Among untreated patients, the mortality rate can reach 20% (20). Human-to-human transmission of R. prowazekii has not been reported. As a biothreat agent, R. prowazekii would likely be spread through the aerosol route or through infected lice. Additional information regarding R. prowazekii can be found in chapter 64.

Transmission and Disease C. perfringens is a spore-forming, Gram-positive bacillus that naturally inhabits the soil. C. perfringens produces several toxins, including epsilon toxin, which is synthesized by toxinotypes B and D. C. perfringens toxinotypes B and D are animal pathogens. Epsilon toxin forms pores in the plasma membranes of eukaryotic cells, but its mode of action is not fully elucidated. In animals, high levels of epsilon toxin are produced in the intestine. The toxin then spreads via the circulatory system to various organs,

Viral Encephalitis (Alphaviruses) Significance Among the alphaviruses, Eastern equine encephalitis (EEE) virus, Venezuelan equine encephalitis (VEE) virus, and Western equine encephalitis (WEE) virus share several characteristics related to their joint classification as potential bioterrorism agents: route of transmission, morbidity in humans and equines, and cell culture infectivity.

224 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

Transmission and Disease The genus Alphavirus is a member of the family Togaviridae. There are 29 species in the genus Alphavirus, including EEE, VEE, and WEE viruses (21). Alphaviruses infecting humans are arthropod borne and cause encephalitis in humans. Diseases can range from mild fever and myalgias to fatal encephalitis. Survivors of encephalitis often suffer from permanent neurologic sequelae. As biothreat agents, alphaviruses would likely be spread through the aerosol route or through infected mosquitoes. Additional information regarding alphaviruses can be found in section IV of this Manual.

Food- and Water-Safety Threats (Bacteria, Viruses, and Protozoa) Significance Contamination of food and water supplies is a real as well as potential route for the dispersal of bioterrorism agents because of the ability to infect a large number of individuals. Food- and waterborne agents considered to be potential bioterrorism agents include Salmonella, Shigella, toxigenic strains of Escherichia coli, Campylobacter, Vibrio cholerae, and Vibrio parahaemolyticus, Listeria monocytogenes, Yersinia enterocolitica, noroviruses, hepatitis A virus, Cryptosporidium parvum, Cyclospora cayetanensis, Entamoeba histolytica, Giardia duodenalis (formerly Giardia lamblia), microsporidia, and Toxoplasma gondii (22). At least two of these agents have been used in documented cases of intentional contamination and bioterrorism. In one event, pastries were intentionally contaminated with a laboratory stock culture of Shigella dysenteriae, infecting 12 (27%) of 45 laboratory staff members (23). In another event, members of a religious commune intentionally contaminated salad bars in several restaurants with Salmonella (24).

Transmission and Disease The food- and waterborne agents listed above share some features, such as the fecal-oral route of transmission (especially secondary cases), yet they also have unique characteristics. Characteristics that are compatible with bioterror use include a low infectious dose (Shigella, norovirus, hepatitis A virus, and C. parvum) and environmental stability (norovirus, hepatitis A virus, and C. parvum), including in aerosols (norovirus). While most of these agents cause an acute or protracted gastroenteritis, some are associated with additional syndromes, such as septicemia (Salmonella enterica serotype Typhi and L. monocytogenes), Guillain-Barré syndrome (Campylobacter jejuni), and hemolytic-uremic syndrome (E. coli serotype O157:H7). Some cause significant morbidity and mortality in immunocompromised persons, including those with AIDS (L. monocytogenes, microsporidia, T. gondii, and C. parvum). A challenge with the prompt identification of bioterrorism events involving food- and waterborne agents is the tendency of these agents to produce large outbreaks of disease naturally. Epidemiological evidence may be necessary to indicate suspicion of bioterrorism. In summary, there is a very real possibility that the laboratory will be the first to recognize an act of bioterrorism. In a suspected or confirmed bioterrorism event, immediate and effective communication with all appropriate institutional and medical personnel and public health officials is imperative. The laboratory may be called upon to assist in the diagnosis and management of patients

who have been exposed overtly or covertly to a bioterrorism agent. The laboratory needs to promptly assist clinicians by providing them with accurate information on the selection, collection, and transport of specimens. In addition, the laboratory must handle these specimens in a manner that will result in the greatest probability of success in establishing a diagnosis and minimizing the exposure of hospital personnel and patients to infectious agents.

REFERENCES 1. Christopher GW, Cieslak TJ, Pavlin JA, Litzen EM. 1997. Biological warfare: a historical perspective. JAMA 278:412–417. 2. Martin GJ, Friedlander AM. 2010. Anthrax as an agent of bioterrorism, p 3983–3992. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 3. Pavani R, Bleck TP. 2010. Botulinum toxin as a biological weapon, p 3993–3994. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 4. Derbes VJ. 1966. De Mussis and the great plague of 1348. A forgotten episode of bacteriological warfare. JAMA 196:59–62. 5. Borio LL, Hynes NA. 2010. Plague as a bioterrorism weapon, p 3965–3970. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 6. Tucker JB. 2000. Toxic Terror. BCSIA Studies in International Security. MIT Press, Cambridge, MA. 7. Rotz LD, Cono J, Damon I. 2010. Smallpox as an agent of bioterrorism, p 3977–3981. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 8. Davis CJ. 1999. Nuclear blindness: an overview of the biological weapons programs of the former Soviet Union and Iraq. Emerg Infect Dis 5:509–512. 9. Gilchrist MJR, McKinney WP, Miller JM, Snyder JW. 2000. Cumitech 33, Laboratory safety, management, and diagnosis of biological agents associated with bioterrorism. Coordinating ed, Snyder JW. ASM Press, Washington, DC. 10. Hodges LS, Penn RL. 2010. Francisella tularensis (tularemia) as an agent of bioterrorism, p 3971–3975. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 11. Peters CJ. 2010. Viral hemorrhagic fevers as agents of bioterrorism, p 3995–3998. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 12. Young EJ. 2010. Brucella species, p 2921–2925. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 13. USAMRIID. 2004. Medical Management of Biological Casualties Handbook, 5th ed. U.S. Army Medical Research Institute of Infectious Diseases, Fort Detrick, Frederick, MD. 14. Marrie TJ, Roult D. 2010. Coxiella burnetii (Q fever), p 2511–2519. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA. 15. Que Y, Moreillon P. 2010. Staphylococcus aureus (including staphylococcal toxin shock), p 2543–2578. In Mandell GL, Bennett JE, Dolin R (ed), Principles and Practices of Infectious Diseases, 7th ed. Elsevier Churchill Livingstone, Philadelphia, PA.

14. Biothreat Agents n 16. Popoff MR. 2011. Epsilon toxin: a fascinating pore-forming toxin. FEBS J 278:4602–4615. 17. Titball RW. 2009. Clostridium perfringens vaccines. Vaccine 27:D44–D47. 18. Beeckman DSA, Vanrompay DCG. 2009. Zoonotic Chlamydophila psittaci infections from a clinical perspective. Clin Microbiol Infect 15:11–17. 19. Walker DH. 2009. The realities of biodefense vaccines against Rickettsia. Vaccine 27:D52–D55. 20. Azad AF. 2007. Pathogenic Rickettsiae as bioterrorism agents. Clin Infect Dis 45:S52–S55. 21. Gould EA, Coutard B, Malet H, Morin B, Jamal S, Weaver S, Gorbalenya A, Moureau G, Baronti C, Delogu I, Forrester N, Khasnatinov M, Gritsun T, de Lamballerie X, Canard B. 2010. Understanding the alphaviruses: recent

225

research on important emerging pathogens and progress towards their control. Antiviral Res 87:111–124. 22. Pappas G, Panagopoulou P, Christou L, Akritidis N. 2006. Category B potential bioterrorism agents: bacteria, viruses, toxins, and foodborne and waterborne pathogens. Infect Dis Clin N Am 20:395–421. 23. Kolavic SA, Kimura A, Simons SL, Slutsker L, Barth S, Haley CE. 1997. An outbreak of Shigella dysenteriae type 2 among laboratory workers due to intentional food contamination. JAMA 278:396–398. 24. Török TJ, Tauxe RV, Wise RP, Livengood JR, Sokolow R, Mauvais S, Birkness KA, Skeels MR, Horan JM, Foster LR. 1997. A large community outbreak of salmonellosis caused by intentional contamination of restaurant salad bars. JAMA 278:389–395.

The Human Microbiome JAMES VERSALOVIC, SARAH K. HIGHLANDER, AND JOSEPH F. PETROSINO

15 Louis, MO). Further efforts were made to standardize how samples were processed, sequenced, and analyzed (4). The progress within the HMP culminated in the publication of two flagship manuscripts (5, 6) and more than 30 companion papers (e.g., see http://www.ploscollections.org/hmp), beginning in June 2012, that describe the initial observations associated with the HMP healthy cohort and numerous derivative studies that examine specific facets of these more deeply characterized communities (Fig. 1). Among initial microbiome comparisons, analyses of samples across 18 body sites confirmed high interindividual variation (7) and determined that even rare organisms in these communities are important reservoirs of genetic diversity (8). Additionally, data from the large HMP cohort suggest that the composition of the gut microbiome is most often characterized by smooth abundance gradients of key organisms (6) and does not necessarily cluster subjects into discrete types. However, microbial communities found in other niches, such as the vagina, can exhibit such clustering (4). While the microbial communities varied among subjects, the metabolic pathways encoded by these organisms and necessary for human-commensal viability were consistently present, forming a functional “core” to the microbiome at all body sites (6, 9, 10). Although the pathways and processes of this core were consistent, the specific genes associated with these pathways varied. Microbial sugar utilization, for example, was enriched for metabolism of simple sugars in the oral cavity, complex carbohydrates in the gut, and glycogen/ peptidoglycan degradation in the vaginal microbiome (10).

The total number of bacteria in the human body is estimated to be at least 10 times greater than the number of human cells (1, 2), and recent studies, particularly the efforts of the NIH Human Microbiome Project (HMP) Consortium, have led to a greater understanding of the identity and distribution of microorganisms that constitute these microbial populations. In particular, implementation of next-generation sequencing (NGS) has helped to illuminate how these bacteria contribute to and are affected by human health and disease. Significant progress in cataloging and characterizing these microorganisms and genes has been made in recent years, thanks to NGS approaches, and microbiome studies have expanded beyond the gut to other body sites. NGS DNA sequencing platforms have made it possible to sequence the DNA of the collective genome (or metagenome) of entire microbial communities (Fig. 1) from different body sites in health and disease and in different life stages, effectively enabling the characterization of the “human microbiome” (see Table 1 for basic definitions).

INTRODUCTION TO THE HUMAN MICROBIOME PROJECT In 2007, an NIH Roadmap for Medical Research project called the Human Microbiome Project (HMP) was initiated (http://nihroadmap.nih.gov/hmp/). The overarching goal of the HMP is to develop tools and resources for characterization of the human microbiota and to relate this microbiota to human health and disease. The HMP is leveraging the constantly advancing sequencing and bioinformatic technologies to address the following broad goals:

TECHNIQUES FOR THE STUDY OF THE HUMAN MICROBIOME

• Determining whether individuals share a core human microbiome • Understanding whether changes in the human microbiome can be correlated with changes in human health and disease • Developing the new technological and bioinformatic tools needed to support these goals • Addressing the ethical, legal, and social implications raised by human microbiome research

Early studies of the human microbiome relied on culturedependent methods; however, it is now known that the majority of microorganisms from the human body cannot be cultured in vitro. Most current techniques for characterization of a metagenomic sample are PCR based and target the highly conserved bacterial small-subunit 16S rRNA gene. Portions of the gene can be amplified and fingerprinted by using electrophoretic techniques, such as terminal restriction fragment length polymorphism (TRFLP) analysis and denaturing gradient gel electrophoresis. Full-length 16S rRNA genes or segments of these genes can be amplified prior to microarray analyses or DNA sequencing studies on

A comprehensive clinical protocol outlined standardized sampling (3) and nucleic acid extraction procedures for the two clinical sampling centers (one at Baylor College of Medicine [BCM] and one at Washington University, St.

doi:10.1128/9781555817381.ch15

226

15. The Human Microbiome n

227

FIGURE 1 Model for microbiome data generation and analyses. (A) Various components (green) and processes (lavender) involved in microbiome projects. (B) Processes of sequencing data generation/editing and data analyses (bioinformatic strategies). Adapted from reference 128 with permission from the Public Library of Science. doi:10.1128/9781555817381.ch15.f1

NGS platforms (e.g., the 454 platform and, more recently, the Illumina MiSeq and Life Technologies Ion Torrent platforms). 16S rRNA gene sequences that are ≥97% identical are considered to be within the same species, while those that are ≥95% identical are within the same genus. Culture-dependent and culture-independent surveys have shown that the human body hosts four predominant phyla (Table 2). The four phyla, Actinobacteria, Bacteroidetes, Firmicutes, and Proteobacteria, are the most highly represented phyla across all body sites (Fig. 2). In a recent study, these four phyla comprised 92.3% of bacterial DNA sequences analyzed from multiple human sources, including hair, oral cavity, skin, and gastrointestinal (GI) tract (11). The predominant phyla vary by anatomical site, and presumably the host milieu has a crucial role in shaping the composition of microbial communities at each site. Recent efforts have been aimed at expanding the DNA sequence representation within each phylum so that more comprehensive phylogenetic assessments can be performed in the future (12).

HUMAN MICROBIOME STUDIES The Oral Microbiome Each human body site is characterized by distinctive patterns of microbial composition (Fig. 2). The oral cavity includes ecologic niches rich in microbial diversity, such as saliva, gingival crevices, the tongue surfaces, and the posterior pharynx, and is colonized with hundreds of bacterial species. An estimated 500 or more different species reside in the human oral cavity (8, 13). Comprehensive sampling of the oral cavity (nine distinct sites), including plaque and saliva, was included in the HMP (5, 6). The human oral microbiome matches or exceeds the intestinal microbiome, and those of all other body sites sampled in the HMP, in terms of microbial diversity within individuals (alpha diversity) (6). The HMP has estimated the presence of approximately 70 distinct bacterial genera in the oral cavities of most adults (6). The most abundant oral bacterial genera include Actinomyces, Bacteroides, Prevotella, Streptococcus, Fusobacterium, Leptotrichia, Corynebacterium, Veillonella,

TABLE 1 Basic terminology of the human microbiome Name Microbiome Metagenome Metatranscriptome Metabolome/metabonome Alpha and beta diversity Operational taxonomic unit Canonical pathogens

Terminology Assemblage of microbes (archaea, bacteria, fungi, parasites, and viruses/phages), constituting a microbial community and occupying a specific habitat. Collection of genomic DNA (genetic capacity) of an entire microbiome and its host DNA. The genes expressed within a microbiome. Metabolites produced by individual microbes (metabolome) or the entire microbiome (metabonome). Microbial species diversity within single samples or human individuals (alpha diversity) and between multiple samples or individuals (beta diversity) at a single body site. Bacterial taxon defined by its 16S rRNA gene sequence or whole-genome sequence. The taxon may or may not have a formally designated species name. Pathogens that cause infections as single infectious agents in a healthy human host.

228 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS TABLE 2

Predominant phyla in healthy individuals by body site

Body site Colon (stool) Esophagus Oral cavity Placenta Skin, dry Skin, moist Skin, sebaceous Small intestine Stomach Upper respiratory tract Vagina

Predominant phylum or phylaa

Reference(s)

Bacteroidetes, Firmicutes, Actinobacteria Firmicutes, Bacteroidetes, Actinobacteria Firmicutes, Actinobacteria, Bacteroidetes Proteobacteria, Tenericutes Proteobacteria, Actinobacteria, Bacteroidetes Actinobacteria, Proteobacteria, Firmicutes, Bacteroidetes Actinobacteria, Firmicutes, Proteobacteria Firmicutes Proteobacteria, Firmicutes, Actinobacteria Firmicutes, Proteobacteria, Actinobacteria

6 44 6, 126 85 110 110 110 2, 53 51, 52 6, 95

Firmicutes, Actinobacteria, Fusobacteria

6, 88, 127

a

Phyla are listed in order of predominance at each body site, based on the cited reference(s).

Rothia, Capnocytophaga, Selenomonas, Treponema, and TM7. Additionally, Methanobrevibacter taxa from the Archaea domain are present in the human oral cavity. Different sites within the oral cavity have various degrees of biological diversity, with plaque and saliva specimens containing the greatest relative diversity (8). New concepts have emerged because of these pioneering studies, including relative differences in biological diversity between body habitats. Relative to human skin, several sites

in the human oral cavity demonstrated relatively greater richness in terms of microbial ecology within individuals but lower beta diversity or a greater similarity of microbial communities between individuals (5, 6). Health-associated microbial communities may protect against infection, but the oral cavity also harbors organisms that are implicated in both local and systemic diseases, including periodontal diseases (14), endocarditis (15), and aspiration pneumonia (16). The relative preponderance of health- or disease-associated mi-

FIGURE 2 Differences in microbial composition by body habitat. The figure shows the relative abundances of six different bacterial phyla at eight different body sites. The relative abundances were obtained by next-generation DNA sequencing of 16S rRNA genes. Adapted from reference 129 with permission from the Nature Publishing Group. doi:10.1128/9781555817381.ch15.f2

15. The Human Microbiome n

crobes combined with human genetic susceptibilities may ultimately account for different disease phenotypes. Periodontitis is an infectious and inflammatory disease condition that is associated with disturbances in microbial ecology among human adult tooth-borne microbiomes. Socransky et al. proposed that periodontitis is the result of complexes or consortia of pathogens (17). The so-called red complex, containing Porphyromonas gingivalis, Tannerella forsythia, and Treponema denticola, is the pathogenic group most strongly associated with this disease process (17, 18). Periodontitis is an independent predictor and comorbidity factor contributing to different disease conditions, including preterm birth, cardiovascular disease, pulmonary disorders, diabetes, and obesity (19). Progressive periodontitis in pregnant women has been reported to increase the risk of severely adverse pregnancy outcomes (20, 21). These associations underscore the significance of the oral microbiome to the systemic health status and predisposition to specific diseases. Microbial community profiling by 16S rRNA gene sequencing and other methods has been used to evaluate the oral microbiome. These methods include denaturing gradient gel electrophoresis (22–24), TRFLP analysis (25– 28), and checkerboard DNA-DNA hybridization, where 45 DNA samples can be queried against 30 to 40 DNA probes (29). Socransky and coworkers applied checkerboard DNADNA hybridization to examine microbial communities in supragingival (30) and subgingival (17) plaque and microbial succession patterns (shifts) during dental biofilm redevelopment in health and periodontitis (31, 32). In both studies, distinct complexes were identified by principal component and correspondence analyses and were assigned to color groups. The communities in supragingival plaque samples from 187 subjects (4,475 samples in total) clustered into six groups, including the aforementioned pathogenassociated red complex. One bacterial pathogen, Aggregatibacter actinomycetemcomitans, has been linked to a highly aggressive periodontitis (localized aggressive periodontitis) in Africans (19). The uncultured bacterial taxon TM7 has been associated with periodontitis based on whole-genome shotgun (WGS) sequencing studies, but this disease association has yet to be evaluated fully in a population-based cohort. As in the HMP, other studies have examined microbial colonization and succession at different sites in the oral cavity. Aas et al. (33) examined nine oral sites: tongue dorsum, tongue lateral sides, buccal epithelium, hard palate, soft palate, supragingival plaque, subgingival plaque, maxillary anterior vestibule, and tonsils. A total of 2,589 16S rRNA gene clones from five subjects were evaluated; this sequencing effort resulted in 141 predominant species and 13 new phylotypes. All sites contained the Gemella, Granulicatella, Streptococcus, and Veillonella genera. The tonsils were found to have the greatest microbial diversity and varied significantly among subjects. Organisms implicated in bacterial endocarditis, such as Streptococcus mitis, Streptococcus oralis (34), and Granulicatella adiacens (35), were found in each healthy subject, which supports the concept that such oral microbes may potentially translocate into the bloodstream secondary to dental procedures. A central concept that emerged from the HMP was that opportunistic pathogens were much more prevalent than canonical pathogens and were present across multiple body sites in a given individual (6). Individuals with breaches in mucosal integrity or immunocompromised status may be vulnerable to opportunistic pathogens present in the human microbiome.

229

A human oral microbe identification microarray (http:// mim.forsyth.org) that contains 16S rRNA gene oligonucleotide probes representing approximately 300 oral bacterial species was described previously (36). While the presence of more periodontal pathogens was observed in the diseased subjects, the main conclusion from the study was that increased microbial diversity was associated with diseased subgingivae. The salivary microbiome has also been studied (37, 38). Samples from 120 healthy subjects from 12 geographic locations were analyzed by Sanger sequencing of the V4 to V6 regions of the 16S rRNA gene. A total of 101 genera were identified, and the numbers of bacterial genera per subject ranged from 6 to 30. The most prevalent genus was Streptococcus (23.7%), consistent with the predominance of Streptococcus in the esophagus and stomach of healthy individuals.

The Gastrointestinal Microbiome The human GI tract encompasses numerous different anatomical sites, including the esophagus, stomach, small intestine, colon, rectum, and anus. Each of these sites is colonized by different numbers and populations of microbes.

Esophageal Microbiome The esophagus is colonized by bacteria that are introduced from the oropharynx by swallowing or from the stomach by reflux. Early studies focused on surveys of cultured bacteria (39–43). Limited numbers of bacteria were examined, and in nondiseased subjects, aerobic and anaerobic Grampositive organisms predominated. Only three metagenomic surveys of the esophageal microbiota have been reported. A study by Pei et al. (44) showed that the distal esophageal microbiomes of four adults had compositions similar to that of the oropharynx, with the exception that no spirochetes were found in the esophagus. As in the oral cavity, Streptococcus was the dominant genus in the healthy esophageal microbiome. Bacterial DNA sequences were categorized into the following six phyla or groups: Firmicutes (70%), Bacteroidetes (20%), Actinobacteria (4%), Proteobacteria (2%), Fusobacteria (2%), and TM7 (1%). This distribution highlights the preponderance of Firmicutes and Bacteroidetes in the GI tract. Thirty-six new species were discovered, and Chao 1 (45) analysis estimated that the esophageal community contains about 140 species-level operational taxonomic units (OTUs) (44). The same group examined differences in the esophageal microbial communities in patients with esophagitis or Barrett’s esophagus (intestinal metaplasia) (46, 47). Two distinct microbiomes were seen in healthy and diseased subjects. Healthy individuals had esophageal microbiomes that were composed predominantly of streptococci, while the diseased patients’ microbiomes had larger numbers of Gram-negative anaerobes and increased bacterial diversity (47). A similar shift in terms of bacterial composition was reported in an earlier study of esophageal carcinoma (41).

Gastric Microbiome The low pH and rapid peristalsis in the stomach suppress persistent colonization by many bacteria. The stomach and small intestine each are thought to contain about 100 microorganisms/ml, but the organismal counts can increase to 105 per ml following a meal (48). The best-studied and most dominant member of the stomach microbiota is Helicobacter pylori (49). Culture-dependent methods have revealed other genera, such as Lactobacillus, Streptococcus, and Staphylococcus, as well as members of the Enterobacteriaceae, in the stomach (50), although a metagenomic analysis of gastric

230 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

biopsy specimens revealed far greater diversity (128 phylotypes) than had been appreciated previously by culturebased approaches (51). The most prevalent genera in the gastric microbiome, besides H. pylori, include several genera found commonly in the oral cavity and esophagus. Bacterial genera found in the human stomach include Prevotella, Streptococcus, Veillonella, and Rothia. The genus Streptococcus is a predominant genus in the stomach when H. pylori is absent (52).

Small Intestinal Microbiome Like the stomach, the proximal small intestine (duodenum and jejunum) is colonized by relatively limited numbers of bacterial species. Acidic pH and bile in the proximal to mid-small intestine inhibit bacterial colonization. Bacterial communities in the small intestine typically consist of lactobacilli, enterococci, Gram-positive aerobes, and facultative anaerobes, and the genus Streptococcus seems to be the dominant genus in the duodenum and jejunum (2, 50, 53). This portion of the GI tract has been assessed mainly by culturedependent methods, with a relative paucity of molecular data. A recent quantitative-PCR-based study described differences in microbial composition between ileostomy specimens and intact small intestinal tissue (54). In the ileum, the microbial composition becomes complex and approaches that of the colon in terms of species richness and the nature of predominant bacterial genera.

Colonic Microbiome The human colon contains a rich collection of microbial communities, with numerical estimates ranging from 1011 to 1012 bacteria/g, and likely exceeding 1,000 bacterial species (6, 8) per individual, the majority of which belong to the phyla Bacteroidetes and Firmicutes (6, 55, 56). Other phyla with sufficient representation in the intestine include Actinobacteria, Proteobacteria, and Verrucomicrobia. In patients with different GI diseases, bacterial population shifts occur such that Proteobacteria become relatively more abundant (57). Comprehensive evaluation of self-collected stool specimens from healthy adults documented high degrees of species richness and microbial diversity in the human intestine (5, 6) relative to other body habitats. The intestine is characterized by a long “tail” of rare, unknown bacterial taxa that accounts for the majority of species or OTUs in the gastrointestinal tract (8). In contrast, in the absence of nucleic acid amplification, the healthy intestinal microbiome contains 70 to 100 known bacterial taxa, on average, based on whole-genome shotgun sequencing (6, 58). In terms of known gut bacteria, the dominant genus by relative abundance in stool is Bacteroides. A relatively abundant signature species is another organism from the phylum Bacteroidetes, known as Prevotella copri (6), and these signature organisms vary between different body habitats. The spatial distribution of gut microbes also depends on the mucosal versus luminal locations of these communities (56). The mucosa-associated microbiomes differ in composition from those found in stool specimens. Bacteria from the distal gut are critical to host nutrition and may play key roles in health and disease. Microbederived carbohydrate fermentation by-products, such as short-chain fatty acids (SCFAs) (e.g., butyrate, acetate, and propionate), provide 10% or more of the body’s metabolic requirements (59). Butyrate, produced by clostridial clusters IV and XIVa, is the primary energy source of the colonic epithelium, and this SCFA has been reported to possess anticancer features (60–62). SCFAs may play a role in preventing infection by pathogens such as Salmonella enterica

serovar Typhimurium (63) and enterohemorrhagic Escherichia coli O157:H7 (64). Colonic bacteria further contribute to nutrition by synthesizing amino acids (65) and vitamins (e.g., vitamins K and B12, biotin, folic acid, and pantothenate) (66, 67). Three basic enterotypes or classes of gut microbiomes have been described, with relatively distinct features in terms of bacterial composition and function (68). Enterotypes 1, 2, and 3 are enriched for different genera, such as Bacteroides, Prevotella, and Ruminococcus, respectively. The same enterotypes are differentially enriched for specific pathways, such as biotin and thiamine biosynthesis (68), and these genetic profiles highlight differences in vitamin and metabolite production by the microbiome. This classification system may be a useful framework for considering basic differences in human populations and susceptibility to disease. For example, differences in biosynthetic capacity, including vitamin metabolism, may have nutritional implications for human health. It is worth noting that the concept of enterotypes in healthy humans has been challenged by alternative concepts, such as that of enterogradients (55). Gut microbial communities may undergo rapid shifts in composition due to dietary changes (69), but enterotypes remained relatively stable despite a 10-day dietary intervention (70). WGS sequence reads from fecal samples of two individuals were analyzed in depth (71). More than 50,000 open reading frames were predicted and examined for enrichment of COG (clusters of orthologous groups) and KEGG (Kyoto Encyclopedia of Genes and Genomes) pathways. Analysis of these pathways revealed enzymes required for degradation of plant polysaccharides, production of SCFAs, vitamin biosynthesis, methanogenesis, and degradation of toxic plant phenolics. These results and those of other studies highlight the metabolic capabilities of the gut microbiota. The gut microbiome appears to have a fundamental impact on human body metabolism, and several studies have highlighted differences in gut microbial populations in obesity and metabolic syndrome. Ley et al. compared the microbiomes in stool samples from obese human subjects placed on fat-restricted and carbohydrate-restricted diets (72). Regardless of the diet type, the percentage of Bacteroidetes organisms increased with weight loss, while the percentage of Firmicutes organisms decreased. A different approach is to examine the relative numbers of different genes in the entire intestinal microbiome or microbial gene richness as a biomarker of human health. Using this approach, individuals have been classified as either high-gene-count (HGC) or low-gene-count (LGC) individuals, and these gene counts seem to affect health status and disease susceptibility. LGC and HGC individuals yielded average quantities of 380,000 and 640,000 bacterial/phage genes, respectively (73). HGC individuals were proposed to have a greater variety of microbial metabolic functions, a relatively robust gut microbiome, and better overall health status, including a lower prevalence of obesity and metabolic disorders (73). Metagenomic studies aimed at understanding the microbial contributions to chronic immune-mediated disorders, such as inflammatory bowel disease (IBD) and type 2 diabetes, have been reported. IBD is likely to be multifactorial in pathogenesis, with factors including host genetics, immune responses, environmental factors such as diet, and the gut microbiome (74). Crohn’s disease and ulcerative colitis are associated with diminished fecal diversity. A PCR-based survey indicated that biopsy samples from patients with IBD were depleted of commensal bacteria belonging to the Firmicutes and Bacteroidetes phyla (75). In particular,

15. The Human Microbiome n

Clostridiales members were diminished in relative abundance, including many bacterial taxa in clostridial cluster XIVa, which contains butyrate-producing organisms. Subsequent studies revealed deficiencies of Faecalibacterium prausnitzii in patients with IBD (75–78). Beyond studies of microbial composition, differences in the relative abundances of metabolic pathways were found in patients with IBD (79). Oxidative stress pathways were relatively enriched in the IBD microbiome, whereas several microbial metabolic pathways associated with carbohydrate metabolism and amino acid biosynthesis were enriched in healthy subjects. IBD genotype (e.g., NOD2 status) and phenotype have been associated with specific compositional changes in the intestinal microbiome (80). Differences in gut microbial composition have been described for type 2 diabetes, a chronic disorder with autoimmune and metabolic components. Metagenome-wide association studies (MGWAS) have identified genetic signatures in the metagenome which are associated with type 2 diabetes (81). These studies strongly suggest that the microbiota in the gut may contribute to activation of signaling pathways required for the development of human diseases such as diabetes.

The Vaginal Microbiome The vaginal microbiota plays an important role in preventing genital and urinary tract infections. The composition of the vaginal microbiota varies with age, pH, and hormonal levels (82). The vaginal microbiome is dynamic and changes during the menstrual cycle (83) and during pregnancy (84). Interestingly, the placental microbiome that develops during pregnancy has also been described (85), and the microbial composition of the placental microbiome is distinct from that of the vaginal microbiome. The placental microbiome is most similar in metagenomic composition to the oral microbiome. In the vagina, lactobacilli are considered the most prevalent organisms in healthy premenopausal women (86, 87), and these are considered protective for the host based on their proposed role in suppression of pathogen colonization. Such effects may result from mucous adherence by lactobacilli, production of organic acids and reduction of vaginal pH, and production of antimicrobial compounds that prevent pathogen proliferation (88). In addition to the lactobacilli, the predominant cultured vaginal microbes in the clinical microbiology laboratory are Mobiluncus spp., Gardnerella vaginalis, Bacteroides spp., Prevotella spp., and Mycoplasma hominis (89). Microbial community profiling studies have demonstrated that the vaginal microbiota is highly variable, and fluctuations in composition presumably depend on differences in sexual and hygienic practices, in addition to host genetics. In the HMP, three sites in the vagina were studied, and differences were documented among these sites (6). The vagina is relatively limited in terms of microbial diversity within and between individuals. Lactobacillus is the predominant member of the vaginal community in most individuals, but in some cases, vaginal lactobacilli may be undetectable. The microbiota of eight healthy women with three different “grades” of vaginosis was examined by sequencing the V1 to V3 regions of the 16S rRNA gene by the Sanger method (90). The vaginal bacterial composition associated with each of these grades was quite distinct: grade 1 individuals (healthy) were almost exclusively colonized by Lactobacillus crispatus, Lactobacillus gasseri, and Lactobacillus jensenii; grade 2 subjects had Lactobacillus iners, Atopobium vaginae, Prevotella bivia, and Sneathia sanguinegens; and grade 3 subjects were predominantly colonized with A. vaginae or Peptostreptococcus anaerobius. Several studies have associated

231

an altered vaginal microbiota with an increased risk of viral coinfection (91–93). Previous studies showed that the microbial composition of the vaginal microbiota varies among healthy Caucasian women (94). The number and distribution of phylotypes differed among the five subjects. Two females had exclusively or nearly exclusively L. crispatus, one had L. iners as the predominant species, and one was predominantly colonized by A. vaginae. The fifth subject had seven phylotypes, predominantly L. iners, but also significant amounts of A. vaginae, Megasphaera, and Leptotrichia. Like lactobacilli, Leptotrichia and Atopobium are lactic acid producers. Different microbial communities may manifest shared functions. A relatively large study included 396 asymptomatic North American women and documented the presence of five groups based on vaginal microbiome composition (87). Four of the five groups were dominated by different Lactobacillus species, whereas the fifth group contained larger proportions of strict anaerobes. The proportions of each microbiome group varied among four ethnic groups included in this study and highlighted the interrelationships between the host genotype/phenotype and vaginal microbiome composition (87). Such differences in microbial communities may contribute to differences in disease susceptibility.

The Microbiome of the Respiratory Tract The healthy nares and nasopharynx contain streptococci, staphylococci, corynebacteria, Moraxella spp., Neisseria spp. (including Neisseria meningitidis), and Haemophilus (95). The HMP documented the relative predominance of the genera Corynebacterium, Moraxella, Propionibacterium, and Staphylococcus in the anterior nares (6). Potential pathogens often colonize the nares and nasopharynx. Microorganisms associated with inner ear infections of children (e.g., Streptococcus pneumoniae, Haemophilus influenzae, and Moraxella catarrhalis) have been found in the nasopharynx (96). The carriage rate for Staphylococcus aureus has been estimated to be approximately 30%, with methicillin-resistant S. aureus representing 1.5% of the strains isolated (97). Studies of the human microbiome reported that Staphylococcus aureus was a prominent species in the anterior nares (6), and this species represents a possible pathogen in its human host. Changes in airway microbial composition have been associated with different disease states involving the respiratory tract. Microbial diversity was greater in patients with tuberculosis than in healthy controls (98), and a core cystic fibrosis-associated microbiome including seven bacterial genera has been proposed (99). The respiratory tract is relatively understudied in terms of its microbiome, but it is becoming clear that significant shifts in composition can be associated with different disease phenotypes.

The Skin Microbiome The skin microbiome is the first line of defense against infection and plays a role in modulating the inflammatory response. Minutes after birth, colonization of the initially sterile skin habitat begins to occur, as newborns are first colonized with similar, low-diversity microbiomes at multiple body sites (100, 101). Members of the environmental microbiota then begin to colonize different regions of the skin as they acquire distinct moisture, temperature, and glandular characteristics, giving rise to habitats with increasingly diverse microbiotas (102). These skin-associated microbial communities continue to change in puberty, depending on age, gender, and environmental exposures (103–107). Metagenomic studies using 16S rRNA gene sequencing in adults showed that the vast majority of skin (as well as

232 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

FIGURE 3 Bacterial compositions at different skin sites. The relative abundances of different bacterial phyla and genera are depicted in terms of the type of skin site (dry, moist, or sebaceous). The data were generated largely by 16S rRNA gene sequencing of organisms on swab samples obtained from different skin sites. Adapted from reference 130 with permission from Elsevier. doi:10.1128/9781555817381.ch15.f3

gut) commensals are grouped into four dominant phyla: Actinobacteria, Firmicutes, Bacteroidetes, and Proteobacteria. These phyla contain thousands of distinct species (103, 108, 109). For instance, the palm microbiome comprised 4,742 distinct species in 51 healthy subjects, with an average of 158 species colonizing a single palm (103). Surveys of more than 20 different skin sites showed that similar habitats, such as the axillae and the popliteal fossae, share similar microbial compositions (Fig. 3) (110). For instance, in all individuals, Propionibacterium species dominate sebaceous areas, such as the back, forehead, and retroauricular crease (6), whereas Staphylococcus and Corynebacterium species dominate moist areas, such as the axillae (Fig. 3). Unexpectedly, Proteobacteria, typically thought to appear on skin only as gastrointestinal contaminants, were found in dry skin habitats, such as the forearm and leg. Diversity and variance fluctuate in a body-site-dependent manner. For example, the antecubital fossa is characterized by fewer distinct species than those at other skin sites. This relatively low alpha diversity is contrasted with a higher beta diversity in comparing the antecubital fossa communities between individuals (6, 111, 112). The skin microbiota generally protects individuals against colonization by pathogens, but skin microbes may be pathogenic if these organisms penetrate the skin in a susceptible host. For example, Staphylococcus epidermidis is a common skin colonizer. However, this species may cause infections in immunocompromised patients or those with indwelling devices. Conversely, Pseudomonas flu-

orescens is thought to be a protective skin organism, because it produces the polyketide antibiotic mupirocin (113), which is active against Gram-positive bacteria, including methicillin-resistant S. aureus (108–110, 114). The skin microbiome varies substantially between individuals, and colonizing microbes may produce antimicrobial compounds that shape microbial communities.

REFERENCE STRAIN GENOMES A critical component of the HMP has been the generation of genome sequences of microbes and viruses that inhabit the human body to create a catalog of reference genomes (115). This catalog serves as a reference data set for metagenomic analyses. Bacterial genome sequences (draft or higher quality) from nearly 5,000 nonpathogenic strains that inhabit the human body have been deposited in the NCBI database, and nearly 2,000 were contributed by the HMP Consortium. The HMP goal is to contribute 3,000 reference genomes (bacterial, fungal, and viral) to the catalog. Community input was sought to nominate strains and isolates for sequencing, and an effort was made to distribute isolates across the five major body sites, although strains from the gastrointestinal tract and oral cavity are overrepresented. These genomes have been added to the comprehensive human microbiome catalog (http://www.hmpdacc.org/ catalog/).

15. The Human Microbiome n 233

The Roche 454 long-tag, paired-end sequencing strategy, originally used by the HMP Consortium for reference genome sequencing, has been replaced by high-coverage (ca. 100× to 400×), short-read sequencing on the Illumina HiSeq platform. Reads are assembled using various assemblers, such as Velvet, ALLPATHs, or the Celera Assembler. Nearly all genome sequences that are currently being deposited are draft WGS sequences that consist of one to dozens of unordered scaffolds. To address the draft nature of these sequences, a set of nomenclature, criteria, and standards for different levels of genome finishing was established (115, 116). The five levels are as follows: (i) standard draft, (ii) high-quality draft, (iii) improved high-quality draft, (iv) annotation-directed improvement, and (v) finished sequence. The levels range from basic assembled contigs (standard draft) to a gold standard complete genome that is assembled into one contiguous piece (finished sequence), has less than 1 error per 100,000 bp, and has all repetitive sequences (including rRNA operons) correctly ordered and placed. Most HMP reference genomes (ca. 70%) are highquality drafts, defined as having >90% of the genome contained in contigs longer than 500 bp, >90% of the genome covered >5-fold, an average contig length of >5 kb, a contig N50 of >5 kb, and a scaffold N50 of >20 kb. The HMP Consortium evaluated methods for gene finding using algorithms such as Glimmer (117) and GeneMark (118). Gene calling was established based on evidence from BLAST searches against the NCBI’s bacterial nonredundant nucleotide and protein databases and alignments to proteins in the Pfam database (http://pfam.sanger.ac.uk/). An automated annotation pipeline was established using tools such as tRNAscan-SE to identify tRNA genes (119) and RNAmmer (120) and Rfam (121) to predict noncoding RNA genes. Gene product annotation is based on evidence from programs such as InterProScan (122), PSORTb (123), and KEGG (http://www.genome.jp/kegg/). Uniform protein naming follows Enzyme Commission nomenclature and descriptors suggested by the JCVI Prokaryotic Annotation Pipeline (124). The utility of the HMP Reference Genomes Catalog was demonstrated by the Human Microbiome Consortium in a read-mapping exercise. A total of 3.5 Tb of Illumina WGS data were generated from 681 microbiome samples from the HMP cohort (5). Following trimming and filtering of human DNA contamination, these reads were aligned to a carefully curated nonredundant reference genome set that contained 2,265 complete and draft genomes plus plasmid sequences that included the sequences of 800 HMP reference genomes (http://hmpdacc.org/HMREFG). In this case, 57.6% of the 35 billion reads were aligned to a reference genome in the HMREFG data set. Furthermore, 26% of these reads could be aligned to one of 223 select HMP reference genomes (5). This result is encouraging, but the coverage is far less than 100%, indicating the importance of continuing reference genome sequencing.

SUMMARY AND CONCLUSIONS The human microbiome includes a diverse collection of human-associated microbes that represent a small fraction of the total microbial “universe” in other animals, life forms, and the environment. This human microbiome can be defined due to the rapid development of advanced DNA sequencing technologies coupled with bioinformatic strategies. Bioinformatic tools include augmented phylogenetic sequence databases, annotation and genome assembly tools, and refined functional approaches to aggregate sequence

data. Metagenomic strategies are being applied to provide glimpses into the variation in microbial composition and function at different body locations and among different individuals. Future approaches in medical microbiology will be shaped in part by developments in the fields of metagenomics and human microbiome research. The identification of single agents of infection will be supplemented by techniques exploring the relative compositions of microbiomes in the context of different infections and other disease states. Differences in microbial composition that are associated with noninfectious immune-mediated disorders may extend the “reach” of the medical microbiology laboratory into other areas of human medicine in the future. Finally, pathogen discovery efforts (see chapter 16) will be enhanced by new metagenomic strategies, and these studies may uncover single etiologic agents of infections as well as changes in microbial communities at specific body sites that may contribute to infectious diseases. This work was supported in part by research support from the National Institutes of Health (NIH) (grants R01 AT004326, R01 DK065075, U01 CA 170930, UH3 DK 083990, and U54HG004973). We also acknowledge the support of the NIH (grant P30 DK56338) to the Texas Medical Center Digestive Diseases Center.

REFERENCES 1. Luckey TD. 1972. Introduction to intestinal microecology. Am J Clin Nutr 25:1292–1294. 2. Savage DC. 1977. Microbial ecology of the gastrointestinal tract. Annu Rev Microbiol 31:107–133. 3. Aagaard K, Petrosino J, Keitel W, Watson M, Katancik J, Garcia N, Patel S, Cutting M, Madden T, Hamilton H, Harris E, Gevers D, Simone G, McInnes P, Versalovic J. 2013. The Human Microbiome Project strategy for comprehensive sampling of the human microbiome and why it matters. FASEB J 27:1012–1022. 4. Human Microbiome Project Consortium. 2012. Evaluation of 16S rDNA-based community profiling for human microbiome research. PLoS One 7:e39315. 5. Human Microbiome Project Consortium. 2012. A framework for human microbiome research. Nature 486:215– 221. 6. Human Microbiome Project Consortium. 2012. Structure, function and diversity of the healthy human microbiome. Nature 486:207–214. 7. Huse SM, Ye Y, Zhou Y, Fodor AA. 2012. A core human microbiome as viewed through 16S rRNA sequence clusters. PLoS One 7:e34242. 8. Li K, Bihan M, Yooseph S, Methe BA. 2012. Analyses of the microbial diversity across the human microbiome. PLoS One 7:e32118. 9. Abubucker S, Segata N, Goll J, Schubert AM, Izard J, Cantarel BL, Rodriguez-Mueller B, Zucker J, Thiagarajan M, Henrissat B, White O, Kelley ST, Methe B, Schloss PD, Gevers D, Mitreva M, Huttenhower C. 2012. Metabolic reconstruction for metagenomic data and its application to the human microbiome. PLoS Comput Biol 8:e1002358. 10. Cantarel BL, Lombard V, Henrissat B. 2012. Complex carbohydrate utilization by the healthy human microbiome. PLoS One 7:e28742. 11. Costello EK, Lauber CL, Hamady M, Fierer N, Gordon JI, Knight R. 2009. Bacterial community variation in human body habitats across space and time. Science 326:1694–1697. 12. Wu D, Hartman A, Ward N, Eisen JA. 2008. An automated phylogenetic tree-based small subunit rRNA taxonomy and alignment pipeline (STAP). PLoS One 3:e2566.

234 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

13. Paster BJ, Olsen I, Aas JA, Dewhirst FE. 2006. The breadth of bacterial diversity in the human periodontal pocket and other oral sites. Periodontol 2000 42:80–87. 14. Rocas IN, Baumgartner JC, Xia T, Siqueira JF Jr. 2006. Prevalence of selected bacterial named species and uncultivated phylotypes in endodontic abscesses from two geographic locations. J Endod 32:1135–1138. 15. Berbari EF, Cockerill FR 3rd, Steckelberg JM. 1997. Infective endocarditis due to unusual or fastidious microorganisms. Mayo Clin Proc 72:532–542. 16. Scannapieco FA. 1999. Role of oral bacteria in respiratory infection. J Periodontol 70:793–802. 17. Socransky SS, Haffajee AD, Cugini MA, Smith C, Kent RL Jr. 1998. Microbial complexes in subgingival plaque. J Clin Periodontol 25:134–144. 18. Holt SC, Ebersole JL. 2005. Porphyromonas gingivalis, Treponema denticola, and Tannerella forsythia: the “red complex,” a prototype polybacterial pathogenic consortium in periodontitis. Periodontol 2000 38:72–122. 19. Curtis MA, Zenobia C, Darveau RP. 2011. The relationship of the oral microbiota to periodontal health and disease. Cell Host Microbe 10:302–306. 20. Goepfert AR, Jeffcoat MK, Andrews WW, Faye-Petersen O, Cliver SP, Goldenberg RL, Hauth JC. 2004. Periodontal disease and upper genital tract inflammation in early spontaneous preterm birth. Obstet Gynecol 104:777–783. 21. Offenbacher S, Boggess KA, Murtha AP, Jared HL, Lieff S, McKaig RG, Mauriello SM, Moss KL, Beck JD. 2006. Progressive periodontal disease and risk of very preterm delivery. Obstet Gynecol 107:29–36. 22. Li Y, Ku CY, Xu J, Saxena D, Caufield PW. 2005. Survey of oral microbial diversity using PCR-based denaturing gradient gel electrophoresis. J Dent Res 84:559–564. 23. Li Y, Saxena D, Barnes VM, Trivedi HM, Ge Y, Xu T. 2006. Polymerase chain reaction-based denaturing gradient gel electrophoresis in the evaluation of oral microbiota. Oral Microbiol Immunol 21:333–339. 24. Rasiah IA, Wong L, Anderson SA, Sissons CH. 2005. Variation in bacterial DGGE patterns from human saliva: over time, between individuals and in corresponding dental plaque microcosms. Arch Oral Biol 50:779–787. 25. Hommez GM, Verhelst R, Claeys G, Vaneechoutte M, De Moor RJ. 2004. Investigation of the effect of the coronal restoration quality on the composition of the root canal microflora in teeth with apical periodontitis by means of T-RFLP analysis. Int Endod J 37:819–827. 26. Hommez GM, Verhelst R, Vaneechoutte M, Claeys G, Vaneechoutte M,De Moor RJ. 2008. Terminal restriction fragment length polymorphism analysis of the microflora in necrotic teeth of patients irradiated in the head and neck region. J Endod 34:1048–1051. 27. Sakamoto M, Huang Y, Ohnishi M, Umeda M, Ishikawa I, Benno Y. 2004. Changes in oral microbial profiles after periodontal treatment as determined by molecular analysis of 16S rRNA genes. J Med Microbiol 53:563–571. 28. Sakamoto M, Takeuchi Y, Umeda M, Ishikawa I, Benno Y. 2003. Application of terminal RFLP analysis to characterize oral bacterial flora in saliva of healthy subjects and patients with periodontitis. J Med Microbiol 52:79–89. 29. Socransky SS, Smith C, Martin L, Paster BJ, Dewhirst FE, Levin AE. 1994. “Checkerboard” DNA-DNA hybridization. Biotechniques 17:788–792. 30. Haffajee AD, Socransky SS, Patel MR, Song X. 2008. Microbial complexes in supragingival plaque. Oral Microbiol Immunol 23:196–205. 31. Teles FR, Teles RP, Sachdeo A, Uzel NG, Song XQ, Torresyap G, Singh M, Papas A, Haffajee AD, Socransky SS. 2012. Comparison of microbial changes in early redeveloping biofilms on natural teeth and dentures. J Periodontol 83:1139–1148.

32. Uzel NG, Teles FR, Teles RP, Song XQ, Torresyap G, Socransky SS, Haffajee AD. 2011. Microbial shifts during dental biofilm re-development in the absence of oral hygiene in periodontal health and disease. J Clin Periodontol 38:612–620. 33. Aas JA, Paster BJ, Stokes LN, Olsen I, Dewhirst FE. 2005. Defining the normal bacterial flora of the oral cavity. J Clin Microbiol 43:5721–5732. 34. Douglas CW, Heath J, Hampton KK, Preston FE. 1993. Identity of viridans streptococci isolated from cases of infective endocarditis. J Med Microbiol 39:179–182. 35. Woo PC, Fung AM, Lau SK, Chan BY, Chiu SK, Teng JL, Que TL, Yung RW, Yuen KY. 2003. Granulicatella adiacens and Abiotrophia defectiva bacteraemia characterized by 16S rRNA gene sequencing. J Med Microbiol 52:137–140. 36. Colombo AP, Boches SK, Cotton SL, Goodson JM, Kent R, Haffajee AD, Socransky SS, Hasturk H, Van Dyke TE, Dewhirst F, Paster BJ. 2009. Comparisons of subgingival microbial profiles of refractory periodontitis, severe periodontitis, and periodontal health using the human oral microbe identification microarray. J Periodontol 80:1421– 1432. 37. Nasidze I, Li J, Quinque D, Tang K, Stoneking M. 2009. Global diversity in the human salivary microbiome. Genome Res 19:636–643. 38. Nasidze I, Quinque D, Li J, Li M, Tang K, Stoneking M. 2009. Comparative analysis of human saliva microbiome diversity by barcoded pyrosequencing and cloning approaches. Anal Biochem 391:64–68. 39. Finlay IG, Wright PA, Menzies T, McArdle CS. 1982. Microbial flora in carcinoma of oesophagus. Thorax 37:181–184. 40. Gagliardi D, Makihara S, Corsi PR, Viana Ade T, Wiczer MV, Nakakubo S, Mimica LM. 1998. Microbial flora of the normal esophagus. Dis Esophagus 11:248–250. 41. Lau WF, Wong J, Lam KH, Ong GB. 1981. Oesophageal microbial flora in carcinoma of the oesophagus. Aust N Z J Surg 51:52–55. 42. Mannell A, Plant M, Frolich J. 1983. The microflora of the oesophagus. Ann R Coll Surg Engl 65:152–154. 43. Pajecki D, Zilberstein B, dos Santos MA, Ubriaco JA, Quintanilha AG, Cecconello I, Gama-Rodrigues J. 2002. Megaesophagus microbiota: a qualitative and quantitative analysis. J Gastrointest Surg 6:723–729. 44. Pei Z, Bini EJ, Yang L, Zhou M, Francois F, Blaser MJ. 2004. Bacterial biota in the human distal esophagus. Proc Natl Acad Sci USA 101:4250–4255. 45. Chao A. 1987. Estimating the population size for capturerecapture data with unequal catchability. Biometrics 43:783–791. 46. Pei Z, Yang L, Peek RM s, Levine SM, Pride DT, Blaser MJ. 2005. Bacterial biota in reflux esophagitis and Barrett’s esophagus. World J Gastroenterol 11:7277–7283. 47. Yang L, Lu X, Nossa CW, Francois F, Peek RM, Pei Z. 2009. Inflammation and intestinal metaplasia of the distal esophagus are associated with alterations in the microbiome. Gastroenterology 137:588–597. 48. Macfarlane S, Macfarlane GT. 2004. Bacterial diversity in the human gut. Adv Appl Microbiol 54:261–289. 49. Cover TL, Blaser MJ. 2009. Helicobacter pylori in health and disease. Gastroenterology 136:1863–1873. 50. Wilson M. 2008. Bacteriology of Humans: An Ecological Perspective. Blackwell Publishing, Malden, MA. 51. Bik EM, Eckburg PB, Gill SR, Nelson KE, Purdom EA, Francois F, Perez-Perez G, Blaser MJ, Relman DA. 2006. Molecular analysis of the bacterial microbiota in the human stomach. Proc Natl Acad Sci USA 103:732–737. 52. Andersson AF, Lindberg M, Jakobsson H, Backhed F, Nyren P, Engstrand L. 2008. Comparative analysis of

15. The Human Microbiome n

53.

54.

55.

56.

57. 58.

59.

60.

61.

62.

63.

64.

65.

66.

67. 68.

human gut microbiota by barcoded pyrosequencing. PLoS One 3:e2836. Zaidel O, Lin HC. 2003. Uninvited guests: the impact of small intestinal bacterial overgrowth on nutritional status. Pract Gastroenterol 27:27–34. Hartman AL, Lough DM, Barupal DK, Fiehn O, Fishbein T, Zasloff M, Eisen JA. 2009. Human gut microbiome adopts an alternative state following small bowel transplantation. Proc Natl Acad Sci USA 106:17187–17192. Backhed F, Fraser CM, Ringel Y, Sanders ME, Sartor RB, Sherman PM, Versalovic J, Young V, Finlay BB. 2012. Defining a healthy human gut microbiome: current concepts, future directions, and clinical applications. Cell Host Microbe 12:611–622. Eckburg PB, Bik EM, Bernstein CN, Purdom E, Dethlefsen L, Sargent M, Gill SR, Nelson KE, Relman DA. 2005. Diversity of the human intestinal microbial flora. Science 308:1635–1638. Pflughoeft KJ, Versalovic J. 2012. Human microbiome in health and disease. Annu Rev Pathol 7:99–122. Qin J, Li R, Raes J, Arumugam M, Burgdorf KS, Manichanh C, Nielsen T, Pons N, Levenez F, Yamada T, Mende DR, Li J, Xu J, Li S, Li D, Cao J, Wang B, Liang H, Zheng H, Xie Y, Tap J, Lepage P, Bertalan M, Batto JM, Hansen T, Le Paslier D, Linneberg A, Nielsen HB, Pelletier E, Renault P, Sicheritz-Ponten T, Turner K, Zhu H, Yu C, Jian M, Zhou Y, Li Y, Zhang X, Qin N, Yang H, Wang J, Brunak S, Dore J, Guarner F, Kristiansen K, Pedersen O, Parkhill J, Weissenbach J, Bork P, Ehrlich SD. 2010. A human gut microbial gene catalogue established by metagenomic sequencing. Nature 464:59– 65. Macfarlane GT, Cummings JH. 1991. The colonic flora, fermentation, and large bowel digestive function, p 51–92. In Phillips S, Pemberton J, Shorter R (ed), The Large Intestine: Physiology, Pathophysiology and Disease. Raven Press, New York, NY. Archer SY, Meng S, Shei A, Hodin RA. 1998. p21(WAF1) is required for butyrate-mediated growth inhibition of human colon cancer cells. Proc Natl Acad Sci USA 95:6791– 6796. Hinnebusch BF, Meng S, Wu JT, Archer SY, Hodin RA. 2002. The effects of short-chain fatty acids on human colon cancer cell phenotype are associated with histone hyperacetylation. J Nutr 132:1012–1017. McIntyre A, Gibson PR, Young GP. 1993. Butyrate production from dietary fibre and protection against large bowel cancer in a rat model. Gut 34:386–391. Lawhon SD, Maurer R, Suyemoto M, Altier C. 2002. Intestinal short-chain fatty acids alter Salmonella typhimurium invasion gene expression and virulence through BarA/ SirA. Mol Microbiol 46:1451–1464. Fukuda S, Toh H, Hase K, Oshima K, Nakanishi Y, Yoshimura K, Tobe T, Clarke JM, Topping DL, Suzuki T, Taylor TD, Itoh K, Kikuchi J, Morita H, Hattori M, Ohno H. 2011. Bifidobacteria can protect from enteropathogenic infection through production of acetate. Nature 469:543–547. Metges CC. 2000. Contribution of microbial amino acids to amino acid homeostasis of the host. J Nutr 130:1857S– 1864S. Albert MJ, Mathan VI, Baker SJ. 1980. Vitamin B12 synthesis by human small intestinal bacteria. Nature 283:781–782. Hill MJ. 1997. Intestinal flora and endogenous vitamin synthesis. Eur J Cancer Prev 6(Suppl 1):S43–S45. Arumugam M, Raes J, Pelletier E, Le Paslier D, Yamada T, Mende DR, Fernandes GR, Tap J, Bruls T, Batto JM, Bertalan M, Borruel N, Casellas F, Fernandez L, Gautier L, Hansen T, Hattori M, Hayashi T, Kleerebezem M,

69.

70.

71.

72.

73.

74.

75.

76.

77.

235

Kurokawa K, Leclerc M, Levenez F, Manichanh C, Nielsen HB, Nielsen T, Pons N, Poulain J, Qin J, SicheritzPonten T, Tims S, Torrents D, Ugarte E, Zoetendal EG, Wang J, Guarner F, Pedersen O, de Vos WM, Brunak S, Dore J, Antolin M, Artiguenave F, Blottiere HM, Almeida M, Brechot C, Cara C, Chervaux C, Cultrone A, Delorme C, Denariaz G, Dervyn R, Foerstner KU, Friss C, van de Guchte M, Guedon E, Haimet F, Huber W, van Hylckama-Vlieg J, Jamet A, Juste C, Kaci G, Knol J, Lakhdari O, Layec S, Le Roux K, Maguin E, Merieux A, Melo Minardi R, M’Rini C, Muller J, Oozeer R, Parkhill J, Renault P, Rescigno M, Sanchez N, Sunagawa S, Torrejon A, Turner K, Vandemeulebrouck G, Varela E, Winogradsky Y, Zeller G, Weissenbach J, Ehrlich SD, Bork P. 2011. Enterotypes of the human gut microbiome. Nature 473:174–180. David LA, Maurice CF, Carmody RN, Gootenberg DB, Button JE, Wolfe BE, Ling AV, Devlin AS, Varma Y, Fischbach MA, Biddinger SB, Dutton RJ, Turnbaugh PJ. 2014. Diet rapidly and reproducibly alters the human gut microbiome. Nature 505:559–563. Wu GD, Chen J, Hoffmann C, Bittinger K, Chen YY, Keilbaugh SA, Bewtra M, Knights D, Walters WA, Knight R, Sinha R, Gilroy E, Gupta K, Baldassano R, Nessel L, Li H, Bushman FD, Lewis JD. 2011. Linking long-term dietary patterns with gut microbial enterotypes. Science 334:105–108. Gill SR, Pop M, Deboy RT, Eckburg PB, Turnbaugh PJ, Samuel BS, Gordon JI, Relman DA, Fraser-Liggett CM, Nelson KE. 2006. Metagenomic analysis of the human distal gut microbiome. Science 312:1355–1359. Ley RE, Turnbaugh PJ, Klein S, Gordon JI. 2006. Microbial ecology: human gut microbes associated with obesity. Nature 444:1022–1023. Le Chatelier E, Nielsen T, Qin J, Prifti E, Hildebrand F, Falony G, Almeida M, Arumugam M, Batto JM, Kennedy S, Leonard P, Li J, Burgdorf K, Grarup N, Jorgensen T, Brandslund I, Nielsen HB, Juncker AS, Bertalan M, Levenez F, Pons N, Rasmussen S, Sunagawa S, Tap J, Tims S, Zoetendal EG, Brunak S, Clement K, Dore J, Kleerebezem M, Kristiansen K, Renault P, Sicheritz-Ponten T, de Vos WM, Zucker JD, Raes J, Hansen T, Bork P, Wang J, Ehrlich SD, Pedersen O, Guedon E, Delorme C, Layec S, Khaci G, van de Guchte M, Vandemeulebrouck G, Jamet A, Dervyn R, Sanchez N, Maguin E, Haimet F, Winogradski Y, Cultrone A, Leclerc M, Juste C, Blottiere H, Pelletier E, LePaslier D, Artiguenave F, Bruls T, Weissenbach J, Turner K, Parkhill J, Antolin M, Manichanh C, Casellas F, Boruel N, Varela E, Torrejon A, Guarner F, Denariaz G, Derrien M, van Hylckama Vlieg JE, Veiga P, Oozeer R, Knol J, Rescigno M, Brechot C, M’Rini C, Merieux A, Yamada T. 2013. Richness of human gut microbiome correlates with metabolic markers. Nature 500:541–546. Sartor RB. 2006. Mechanisms of disease: pathogenesis of Crohn’s disease and ulcerative colitis. Nat Clin Pract Gastroenterol Hepatol 3:390–407. Frank DN, St Amand AL, Feldman RA, Boedeker EC, Harpaz N, Pace NR. 2007. Molecular-phylogenetic characterization of microbial community imbalances in human inflammatory bowel diseases. Proc Natl Acad Sci USA 104:13780–13785. Mariat D, Firmesse O, Levenez F, Guimaraes V, Sokol H, Dore J, Corthier G, Furet JP. 2009. The Firmicutes/ Bacteroidetes ratio of the human microbiota changes with age. BMC Microbiol 9:123. Martinez-Medina M, Aldeguer X, Gonzalez-Huix F, Acero D, Garcia-Gil LJ. 2006. Abnormal microbiota composition in the ileocolonic mucosa of Crohn’s disease patients as revealed by polymerase chain reaction-denaturing

236 n

78.

79.

80.

81.

82.

83.

84.

85.

86. 87.

88.

89.

90.

91.

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

gradient gel electrophoresis. Inflamm Bowel Dis 12:1136– 1145. Wang W, Chen L, Zhou R, Wang X, Song L, Huang S, Wang G, Xia B. 2014. Increased proportions of Bifidobacterium and the Lactobacillus group and loss of butyrate-producing bacteria in inflammatory bowel disease. J Clin Microbiol 52:398–406. Morgan XC, Tickle TL, Sokol H, Gevers D, Devaney KL, Ward DV, Reyes JA, Shah SA, LeLeiko N, Snapper SB, Bousvaros A, Korzenik J, Sands BE, Xavier RJ, Huttenhower C. 2012. Dysfunction of the intestinal microbiome in inflammatory bowel disease and treatment. Genome Biol 13:R79. Frank DN, Robertson CE, Hamm CM, Kpadeh Z, Zhang T, Chen H, Zhu W, Sartor RB, Boedeker EC, Harpaz N, Pace NR, Li E. 2011. Disease phenotype and genotype are associated with shifts in intestinal-associated microbiota in inflammatory bowel diseases. Inflamm Bowel Dis 17:179–184. Qin J, Li Y, Cai Z, Li S, Zhu J, Zhang F, Liang S, Zhang W, Guan Y, Shen D, Peng Y, Zhang D, Jie Z, Wu W, Qin Y, Xue W, Li J, Han L, Lu D, Wu P, Dai Y, Sun X, Li Z, Tang A, Zhong S, Li X, Chen W, Xu R, Wang M, Feng Q, Gong M, Yu J, Zhang Y, Zhang M, Hansen T, Sanchez G, Raes J, Falony G, Okuda S, Almeida M, LeChatelier E, Renault P, Pons N, Batto JM, Zhang Z, Chen H, Yang R, Zheng W, Yang H, Wang J, Ehrlich SD, Nielsen R, Pedersen O, Kristiansen K. 2012. A metagenome-wide association study of gut microbiota in type 2 diabetes. Nature 490:55–60. Garcia-Closas M, Herrero R, Bratti C, Hildesheim A, Sherman ME, Morera LA, Schiffman M. 1999. Epidemiologic determinants of vaginal pH. Am J Obstet Gynecol 180:1060–1066. Gajer P, Brotman RM, Bai G, Sakamoto J, Schutte UM, Zhong X, Koenig SS, Fu L, Ma ZS, Zhou X, Abdo Z, Forney LJ, Ravel J. 2012. Temporal dynamics of the human vaginal microbiota. Sci Transl Med 4:132ra152. Aagaard K, Riehle K, Ma J, Segata N, Mistretta TA, Coarfa C, Raza S, Rosenbaum S, Van den Veyver I, Milosavljevic A, Gevers D, Huttenhower C, Petrosino J, Versalovic J. 2012. A metagenomic approach to characterization of the vaginal microbiome signature in pregnancy. PLoS One 7:e36466. Aagaard K, Ma J, Antony KM, Ganu R, Petrosino J, Versalovic J. 2014. The placenta harbors a unique microbiome. Sci Transl Med 6:237ra265. Larsen B. 1993. Vaginal flora in health and disease. Clin Obstet Gynecol 36:107–121. Ravel J, Gajer P, Abdo Z, Schneider GM, Koenig SS, McCulle SL, Karlebach S, Gorle R, Russell J, Tacket CO, Brotman RM, Davis CC, Ault K, Peralta L, Forney LJ. 2011. Vaginal microbiome of reproductive-age women. Proc Natl Acad Sci USA 108(Suppl 1):4680–4687. Boris S, Barbes C. 2000. Role played by lactobacilli in controlling the population of vaginal pathogens. Microbes Infect 2:543–546. Holst E, Wathne B, Hovelius B, Mardh PA. 1987. Bacterial vaginosis: microbiological and clinical findings. Eur J Clin Microbiol 6:536–541. Verhelst R, Verstraelen H, Claeys G, Verschraegen G, Delanghe J, Van Simaey L, De Ganck C, Temmerman M, Vaneechoutte M. 2004. Cloning of 16S rRNA genes amplified from normal and disturbed vaginal microflora suggests a strong association between Atopobium vaginae, Gardnerella vaginalis and bacterial vaginosis. BMC Microbiol 4:16. Kaul R, Nagelkerke NJ, Kimani J, Ngugi E, Bwayo JJ, Macdonald KS, Rebbaprgada A, Fonck K, Temmerman M, Ronald AR, Moses S. 2007. Prevalent herpes simplex

92.

93.

94.

95.

96.

97.

98.

99.

100.

101. 102.

103.

104.

105. 106. 107. 108.

109.

virus type 2 infection is associated with altered vaginal flora and an increased susceptibility to multiple sexually transmitted infections. J Infect Dis 196:1692–1697. Myer L, Denny L, Telerant R, Souza M, Wright TC Jr, Kuhn L. 2005. Bacterial vaginosis and susceptibility to HIV infection in South African women: a nested casecontrol study. J Infect Dis 192:1372–1380. Schwebke JR. 2005. Abnormal vaginal flora as a biological risk factor for acquisition of HIV infection and sexually transmitted diseases. J Infect Dis 192:1315–1317. Zhou X, Brown CJ, Abdo Z, Davis CC, Hansmann MA, Joyce P, Foster JA, Forney LJ. 2007. Differences in the composition of vaginal microbial communities found in healthy Caucasian and black women. ISME J 1:121–133. Hull MW, Chow AW. 2007. Indigenous microflora and innate immunity of the head and neck. Infect Dis Clin North Am 21:265–282. Konno M, Baba S, Mikawa H, Hara K, Matsumoto F, Kaga K, Nishimura T, Kobayashi T, Furuya N, Moriyama H, Okamoto Y, Furukawa M, Yamanaka N, Matsushima T, Yoshizawa Y, Kohno S, Kobayashi K, Morikawa A, Koizumi S, Sunakawa K, Inoue M, Ubukata K. 2006. Study of nasopharyngeal bacterial flora. Second report. Variations in nasopharyngeal bacterial flora in children aged 6 years or younger when administered antimicrobial agents. J Infect Chemother 12:305–330. Gorwitz RJ, Kruszon-Moran D, McAllister SK, McQuillan G, McDougal LK, Fosheim GE, Jensen BJ, Killgore G, Tenover FC, Kuehnert MJ. 2008. Changes in the prevalence of nasal colonization with Staphylococcus aureus in the United States, 2001–2004. J Infect Dis 197:1226–1234. Cui Z, Zhou Y, Li H, Zhang Y, Zhang S, Tang S, Guo X. 2012. Complex sputum microbial composition in patients with pulmonary tuberculosis. BMC Microbiol 12:276. van der Gast CJ, Walker AW, Stressmann FA, Rogers GB, Scott P, Daniels TW, Carroll MP, Parkhill J, Bruce KD. 2011. Partitioning core and satellite taxa from within cystic fibrosis lung bacterial communities. ISME J 5:780– 791. Dominguez-Bello MG, Costello EK, Contreras M, Magris M, Hidalgo G, Fierer N, Knight R. 2010. Delivery mode shapes the acquisition and structure of the initial microbiota across multiple body habitats in newborns. Proc Natl Acad Sci USA 107:11971–11975. Sarkany I, Gaylarde CC. 1968. Bacterial colonisation of the skin of the newborn. J Pathol Bacteriol 95:115–122. Capone KA, Dowd SE, Stamatas GN, Nikolovski J. 2011. Diversity of the human skin microbiome early in life. J Invest Dermatol 131:2026–2032. Fierer N, Hamady M, Lauber CL, Knight R. 2008. The influence of sex, handedness, and washing on the diversity of hand surface bacteria. Proc Natl Acad Sci USA 105:17994–17999. Giacomoni PU, Mammone T, Teri M. 2009. Genderlinked differences in human skin. J Dermatol Sci 55:144– 149. Marples RR. 1982. Sex, constancy, and skin bacteria. Arch Dermatol Res 272:317–320. Somerville DA. 1969. The effect of age on the normal bacterial flora of the skin. Br J Dermatol 81(Suppl 1):14+. Somerville DA. 1969. The normal flora of the skin in different age groups. Br J Dermatol 81:248–258. Gao Z, Tseng CH, Pei Z, Blaser MJ. 2007. Molecular analysis of human forearm superficial skin bacterial biota. Proc Natl Acad Sci USA 104:2927–2932. Grice EA, Kong HH, Renaud G, Young AC, Bouffard GG, Blakesley RW, Wolfsberg TG, Turner ML, Segre JA. 2008. A diversity profile of the human skin microbiota. Genome Res 18:1043–1050.

15. The Human Microbiome n 110. Grice EA, Kong HH, Conlan S, Deming CB, Davis J, Young AC, Bouffard GG, Blakesley RW, Murray PR, Green ED, Turner ML, Segre JA. 2009. Topographical and temporal diversity of the human skin microbiome. Science 324:1190–1192. 111. Roth RR, James WD. 1988. Microbial ecology of the skin. Annu Rev Microbiol 42:441–464. 112. Chiller K, Selkin BA, Murakawa GJ. 2001. Skin microflora and bacterial infections of the skin. J Invest Dermatol Symp Proc 6:170–174. 113. Fuller AT, Mellows G, Woolford M, Banks GT, Barrow KD, Chain EB. 1971. Pseudomonic acid: an antibiotic produced by Pseudomonas fluorescens. Nature 234:416– 417. 114. Sutherland R, Boon RJ, Griffin KE, Masters PJ, Slocombe B, White AR. 1985. Antibacterial activity of mupirocin (pseudomonic acid), a new antibiotic for topical use. Antimicrob Agents Chemother 27:495–498. 115. Nelson KE, Weinstock GM, Highlander SK, Worley KC, Creasy HH, Wortman JR, Rusch DB, Mitreva M, Sodergren E, Chinwalla AT, Feldgarden M, Gevers D, Haas BJ, Madupu R, Ward DV, Birren BW, Gibbs RA, Methe B, Petrosino JF, Strausberg RL, Sutton GG, White OR, Wilson RK, Durkin S, Giglio MG, Gujja S, Howarth C, Kodira CD, Kyrpides N, Mehta T, Muzny DM, Pearson M, Pepin K, Pati A, Qin X, Yandava C, Zeng Q, Zhang L, Berlin AM, Chen L, Hepburn TA, Johnson J, McCorrison J, Miller J, Minx P, Nusbaum C, Russ C, Sykes SM, Tomlinson CM, Young S, Warren WC, Badger J, Crabtree J, Markowitz VM, Orvis J, Cree A, Ferriera S, Fulton LL, Fulton RS, Gillis M, Hemphill LD, Joshi V, Kovar C, Torralba M, Wetterstrand KA, Abouellleil A, Wollam AM, Buhay CJ, Ding Y, Dugan S, FitzGerald MG, Holder M, Hostetler J, Clifton SW, Allen-Vercoe E, Earl AM, Farmer CN, Liolios K, Surette MG, Xu Q, Pohl C, WilczekBoney K, Zhu D. 2010. A catalog of reference genomes from the human microbiome. Science 328:994–999. 116. Chain PS, Grafham DV, Fulton RS, Fitzgerald MG, Hostetler J, Muzny D, Ali J, Birren B, Bruce DC, Buhay C, Cole JR, Ding Y, Dugan S, Field D, Garrity GM, Gibbs R, Graves T, Han CS, Harrison SH, Highlander S, Hugenholtz P, Khouri HM, Kodira CD, Kolker E, Kyrpides NC, Lang D, Lapidus A, Malfatti SA, Markowitz V, Metha T, Nelson KE, Parkhill J, Pitluck S, Qin X, Read TD, Schmutz J, Sozhamannan S, Sterk P, Strausberg RL, Sutton G, Thomson NR, Tiedje JM, Weinstock G, Wollam A, Detter JC. 2009. Genomics. Genome project standards in a new era of sequencing. Science 326:236–237. 117. Delcher AL, Harmon D, Kasif S, White O, Salzberg SL. 1999. Improved microbial gene identification with GLIMMER. Nucleic Acids Res 27:4636–4641.

237

118. Borodovsky M, Mills R, Besemer J, Lomsadze A. 2003. Prokaryotic gene prediction using GeneMark and GeneMark.hmm. Curr Protoc Bioinformatics Chapter 4:Unit 4.5. 119. Lowe TM, Eddy SR. 1997. tRNAscan-SE: a program for improved detection of transfer RNA genes in genomic sequence. Nucleic Acids Res 25:955–964. 120. Lagesen K, Hallin P, Rodland EA, Staerfeldt HH, Rognes T, Ussery DW. 2007. RNAmmer: consistent and rapid annotation of ribosomal RNA genes. Nucleic Acids Res 35:3100–3108. 121. Burge SW, Daub J, Eberhardt R, Tate J, Barquist L, Nawrocki EP, Eddy SR, Gardner PP, Bateman A. 2013. Rfam 11.0: 10 years of RNA families. Nucleic Acids Res 41:D226–D232. 122. Quevillon E, Silventoinen V, Pillai S, Harte N, Mulder N, Apweiler R, Lopez R. 2005. InterProScan: protein domains identifier. Nucleic Acids Res 33:W116–W120. 123. Yu NY, Wagner JR, Laird MR, Melli G, Rey S, Lo R, Dao P, Sahinalp SC, Ester M, Foster LJ, Brinkman FS. 2010. PSORTb 3.0: improved protein subcellular localization prediction with refined localization subcategories and predictive capabilities for all prokaryotes. Bioinformatics 26:1608–1615. 124. Davidsen T, Beck E, Ganapathy A, Montgomery R, Zafar N, Yang Q, Madupu R, Goetz P, Galinsky K, White O, Sutton G. 2010. The comprehensive microbial resource. Nucleic Acids Res 38:D340–D345. 125. Tap J, Mondot S, Levenez F, Pelletier E, Caron C, Furet JP, Ugarte E, Munoz-Tamayo R, Paslier DL, Nalin R, Dore J, Leclerc M. 2009. Towards the human intestinal microbiota phylogenetic core. Environ Microbiol 11:2574– 2584. 126. Paster BJ, Boches SK, Galvin JL, Ericson RE, Lau CN, Levanos VA, Sahasrabudhe A, Dewhirst FE. 2001. Bacterial diversity in human subgingival plaque. J Bacteriol 183:3770–3783. 127. Brown CJ, Wong M, Davis CC, Kanti A, Zhou X, Forney LJ. 2007. Preliminary characterization of the normal microbiota of the human vulva using cultivationindependent methods. J Med Microbiol 56:271–276. 128. Gevers D, Knight R, Petrosino JF, Huang K, McGuire AL, Birren BW, Nelson KE, White O, Methe BA, Huttenhower C. 2012. The Human Microbiome Project: a community resource for the healthy human microbiome. PLoS Biol 10:e1001377. 129. Cho I, Blaser MJ. 2012. The human microbiome: at the interface of health and disease. Nat Rev Genet 13:260– 270. 130. Chen YE, Tsao H. 2013. The skin microbiome: current perspectives and future challenges. J Am Acad Dermatol 69:143–155.

Microbial Genomics and Pathogen Discovery* JENNIFER K. SPINLER, PEERA HEMARAJATA, AND JAMES VERSALOVIC

16 In recent years, our understanding of microbial diversity has grown tremendously as many previously unidentified bacterial, archaeal, and viral species have been discovered and sequenced. In the era of the human microbiome and metagenomics (chapter 15), large-scale DNA sequencing projects and advances in bioinformatics have yielded abundant data regarding human-associated microbes. As human microbiology rapidly expands beyond its past framework of cultured pathogens in the medical microbiology laboratory, opportunities for detection and identification of novel human pathogens associated with infectious diseases abound. In this chapter, we focus on specific or defined sets of pathogens associated with human infections, in contrast to microbial components of disease and microbial ecology (topics covered in chapter 15). We begin with an overview of historical methodologies, followed by a brief description of the evolution of nucleic acid sequencing technologies. Finally, we describe how microarrays, nucleic acid sequencing technologies, and mass spectrometry are profoundly reshaping strategies aimed at pathogen discovery and identification. A timeline of these scientific advances is depicted in Fig. 1. Our ability to visualize human microbes first arose in the 17th century when Anton van Leeuwenhoek created the microscope and observed human-associated bacteria in the human oral cavity, which provided the first physical evidence of the diversity and ubiquity of microbes in the world. Another giant leap occurred in the 19th century when Robert Koch first demonstrated that bacteria could be grown in pure culture, beginning with the analysis of blood from cows infected with the anthrax agent. He subsequently became most widely known for the postulates regarding microbial disease causation that bear his name. For decades following, microbiologic culture and microscopy were the only tools available to directly examine human microbes. In the last 2 decades of the 20th century, however, DNA amplification of consensus rRNA gene sequences and other genetic targets revealed a vast diversity of bacteria, viruses, and other microbes that had eluded cultivation in the laboratory. Today, DNA and RNA sequencing technologies have evolved to the point that it is feasible to comprehensively define the collection of microbes present in humans (i.e., the human “microbiome”) or any other ecological

niche in a fashion that is independent of microbiologic culture or microscopy.

A BRIEF HISTORY OF PATHOGEN DISCOVERY Classical Methods Classical methods of microbial discovery relied on the ability to culture microorganisms in the laboratory and to directly visualize microbial morphology by microscopy. Body fluid or tissue specimens associated with specific disease phenotypes and considered to be of infectious origin are still used today to inoculate microbiologic growth media and cultivate the microbe(s) present in the sample. In the case of suspected bacterial agents, selective or nonselective growth media can be utilized, and biochemical or serological tests can facilitate identification of cultured isolates. Primary or immortalized human and mammalian cell lines are inoculated for suspected viruses, and viruses have been identified by cytopathic effects (light microscopy), immunological methods (serological reactions based on different antisera), and viral particle morphology (electron microscopy). For pathogen discovery and biomedical research purposes, clinical specimens have been used for decades to infect animal models and prove Koch’s postulates. These classical methods have been very useful and resulted in the discovery of many currently accepted human pathogens; examples include Bacillus anthracis, Mycobacterium tuberculosis, Yellow fever virus, and Poliovirus. However, there are two fundamental limitations with conventional approaches: (i) these methods depend on the ability of microbes to grow on media substrates in the laboratory; and (ii) even if the microbe can be cultivated, unknown candidate agents may not be unambiguously identified.

Molecular Strategies: Pathogen Discovery In the 1980s, scientists began to apply DNA amplification technologies to detect microbes present in human specimens if existing knowledge about DNA sequences in microbial genomes could be targeted. The recognition that selecting PCR primers designed to highly conserved regions in a set of sequences (e.g., multiple bacteria or several viruses from a common taxonomic group) could enable the detection of previously unidentified microbes provided a novel approach for the identification of microbes. One of the broadest applications of this technique has been the design of primers to

*This chapter contains information presented by Anne M. Gaynor and David Wang in chapter 14 of the 10th edition of this Manual.

doi:10.1128/9781555817381.ch16

238

16. Microbial Genomics and Pathogen Discovery n

239

FIGURE 1 A timeline depicting major landmarks in medical microbiology and molecular microbiology that have enhanced pathogen discovery. doi:10.1128/9781555817381.ch16.f1

the conserved 16S rRNA gene, which enables the detection of most members of the bacterial domain depending on the selection of specific primer pairs. Alternatively, more specific primers can be selected that are conserved within a given taxon (family, genus, or species) to identify a targeted set of microbes. The ability to design highly conserved primers depends on the existence of sufficient amounts of DNA sequencing data, and recent advances in the Human Microbiome Project have contributed greatly to a refinement of primer selection and amplification strategies for

bacterial genera and species. Different microbes have been discovered by using PCR in conjunction with the classical methods of microbial detection. With the advent of Sanger sequencing, entire 16S rRNA genes could be sequenced, including the Escherichia coli 16S rRNA gene in 1978 (1). Relman and coworkers described the first examples of using consensus PCR primers to identify the causative agents of specific human diseases. Bacillary angiomatosis was commonly considered to be infectious in origin. A putative agent could be visualized in tissue sections following staining,

240 n

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

but efforts to culture the organism had failed. Amplicon sequencing using 16S rRNA gene consensus primers demonstrated that a previously uncharacterized rickettsia-like bacterium was present in tissue samples of patients with bacillary angiomatosis (2). The bacterium was identified as a member of the genus Bartonella. A similar approach was subsequently applied to address the etiology of Whipple’s disease. Whipple’s disease was first described in 1907 as a rare systemic disorder that primarily caused malabsorption but could affect any part of the body. Consensus PCR using primers targeting bacterial 16S ribosomal genes resulted in the identification of an uncharacterized actinomycete, which was classified as Tropheryma whipplei (3). These two cases demonstrated the power of molecular pathogen discovery methods. Conserved PCR primers and nucleic acid probes have been applied to virus discovery. However, since no universally conserved sequence akin to 16S rRNA sequences in bacteria is present in all viruses, different consensus nucleic acid sequences must be identified for different classes of viruses and consensus primers designed for each viral taxon of interest. A seminal example of using consensus PCR to identify a viral pathogen occurred during the emergence of hantavirus pulmonary syndrome in 1993 (4). In the course of investigating an unusual outbreak of a lethal pulmonary disease in otherwise healthy young adults in the southwestern United States, extensive testing by classical microbiological methods ruled out the most likely candidates known to cause severe respiratory disease. Serological tests revealed that patient sera were cross-reactive with known hantaviruses. PCR primers were designed to conserved regions of known hantavirus sequences, which were then used to amplify nucleic acids extracted from tissue samples isolated from dying patients. Sequencing of the amplicon generated by the primers resulted in the identification of a novel member of this family, which was ultimately named Sin Nombre virus. Since these seminal applications of consensus PCR for the identification of bacterial and viral pathogens, many applications of nucleic acid amplification for pathogen discovery have been published. Consensus PCR, either alone or in conjunction with microbiologic culture, microscopy, and antigen detection methods, has been used to detect novel microbes, as illustrated by the recent discoveries of new groups of rhinoviruses (5–9), parechoviruses (10–15), Chapare virus (arenavirus) (16), Bundibugyo ebolavirus (17), the Middle East respiratory syndrome coronavirus (18), and an entirely new bacterium associated with cord colitis syndrome (19). However, broad-range PCR strategies often failed to identify a specific infectious agent. In these examples, the authors had robust hypotheses regarding the nature of the microbes of interest (i.e., bacterium versus virus) or which specific candidate viral taxon might be present. Often, leading candidates or phylogenetic groups of interest may not be apparent, limiting the application of specific molecular strategies. Outside of the bacterial kingdom, the relative paucity of consensus probes or primers for archaea, eukaryotic microbes, and viruses emphasizes the limits of targeted molecular strategies. For these reasons, more comprehensive genomics-based strategies have been applied to pathogen discovery and in-depth characterization.

Early Applications of Microbial Genomics: Pathogen Discovery The discoveries of Hepatitis C virus (HCV) and Human herpesvirus 8, also named Kaposi’s sarcoma-associated herpesvirus, represented two breakthroughs in the application

of genomics-based molecular methods for pathogen discovery. In 1989, the identification of HCV in patients with non-A, non-B (NANB) hepatitis relied on a DNA librarybased immunoscreening strategy (20). A randomly primed cDNA library was made from material from infected animals and screened using patient serum from NANB hepatitis patients with the goal of identifying cDNA clones that generated peptide sequences recognized by the patient sera. More than 1 million clones were screened, and a single clone reacted specifically with NANB hepatitis patient sera. From this initial cDNA clone fragment, the entire HCV genome was eventually sequenced. HCV is now recognized as being responsible for the vast majority of cases of NANB hepatitis. In 1994, human herpesvirus 8 was discovered in the lesions of AIDS-associated Kaposi’s sarcoma (21). The identification of Kaposi’s sarcoma-associated herpesvirus relied on representational difference analysis, a subtractive hybridization-based method, to enrich for and identify unique sequences present in Kaposi’s sarcoma lesions but not in healthy tissue controls. While these two examples demonstrated the potential utility of early genomics-based strategies, subsequent success stories were rare, most likely due to the technical challenges associated with first-generation genomics-based strategies. By the end of the 20th century, classical microbiologic culture-based methods for microbial discovery had been augmented by early molecular and genomics-based strategies, such as consensus PCR, library immunoscreening, and representational difference analysis. In parallel, reference genome sequencing of specific microbes was becoming feasible, setting the stage for the convergence of pathogen discovery and microbial genomic sequencing efforts. Further developments in genomics-based strategies were needed to advance pathogen discovery in the 21st century.

RECENT HISTORY OF MICROBIAL GENOMICS Microbe sequencing in the 20th century relied exclusively on Sanger dideoxy sequencing, the dominant sequencing strategy since its invention in 1977. From its initial incarnation using slab-gel electrophoresis, incremental advances in sequencing capacity evolved as sequencing transitioned to capillary electrophoresis. Formally, the era of microbial genomics began with the complete sequencing of the Haemophilus influenzae genome in 1995. However, it was recognized almost 2 decades earlier that an organism’s genomic sequence could serve to classify and define the relatedness of both prokaryotic and eukaryotic organisms (22). When the H. influenzae genome was sequenced, this bacterium became the first free-living organism to have its genome sequenced in its entirety (23). This was a landmark achievement, notable also because of the use of a “shotgun” strategy to assemble the complete genome. “Shotgun” refers to the random fragmentation and cloning of DNA fragments followed by computational assembly of the overlapping regions to generate a complete genome sequence. Based on this proof of principle, genomes of larger microbes and eukaryotic organisms were subsequently sequenced. The following year, Saccharomyces cerevisiae was the first eukaryotic organism to be fully sequenced (24), and then in 1998, the first multicellular eukaryotic genome to be sequenced, that of Caenorhabditis elegans, was published (25). Since then, the complete genomes of many human and animal pathogens have been sequenced, including notable pathogens such as M. tuberculosis (26), Yersinia pestis (27), and Plasmodium falciparum (28). In 2004, the complete 1.2-

16. Microbial Genomics and Pathogen Discovery n 241

Mb genome of mimivirus, the largest known virus, was published (29). Next-generation DNA sequencing modalities have revolutionized the depth and throughput of sequencing applications in medical microbiology. Three major platforms in current use are 454 (454 Life Sciences, a Roche Company, Branford, CT), HiSeq and MiSeq (Illumina, San Diego, CA), and Ion Torrent/PGM (Life Technologies, Carlsbad, CA). Key characteristics of these platforms include the fact that all of them have exponentially increased the raw sequence generation capacity and dramatically decreased the cost per base relative to Sanger sequencing. For a more detailed description of each of these platforms and capabilities, see reference 30. With these increases in sequencing capacity, the sequencing of microbial genomes has become routine. Next-generation DNA sequencing platforms have enabled scientists to comprehensively explore the microbial diversity of the human microbiome and the viral diversity, the “virome,” present in humans (see chapter 15). These efforts have vastly expanded the world of sequenced microbes, and sequences have been deposited in various databases (e.g., http://www.ncbi.nlm.nih.gov/genome/ and http://hmpdacc.org).

THE MERGING OF PATHOGEN DISCOVERY AND MICROBIAL GENOMICS The fields of pathogen discovery and microbial genomics have converged in the past 2 decades so that more pathogens can be detected and identified (Fig. 1). Two basic molecular approaches for massively parallel analysis have recently emerged that are capable of defining the spectrum of microbes present in clinical specimens: microarrays and sequencing-based detection. Both strategies benefited greatly from the increased focus on human and microbial sequencing. Multiple databases of nucleic acid sequences provide the substrate from which consensus PCR primers for sequencing applications and probes for microarrays have been designed. DNA sequencing-based approaches have evolved rapidly in the past decade and have resulted in next-generation DNA sequencing pipelines for pathogen discovery and detection. As sequencing technologies are rapidly supplanting microarray-based hybridization strategies, microarrays continue to represent a useful option for bacterial and viral detection. Data analysis or bioinformatics tools and pipelines are also being refined to meet the challenge of genomics and “big data” in the clinical laboratory, whether the data are generated by microarray or sequencing platforms.

Microarray-Based Approaches Microarrays were initially developed in the early 1990s to analyze gene expression patterns of a single organism in a highly parallel fashion. For example, early studies analyzed every gene present in the yeast S. cerevisiae (31). Regardless of the technical format, the fundamental principle driving all microarrays is that of nucleic acid hybridization. If a given sample contains sequences that are complementary to those represented on the array, hybridization should occur. Thus, the target sequences to be detected by a given microarray are limited only by the availability of DNA sequences from which probes can be designed. As DNA microarray technology evolved in the late 1990s, it became clear that microarrays might make excellent microbial diagnostic tools. The inherently highly parallel nature of microarrays, coupled with the expanding amount of microbial sequencing data, made it possible to design microarrays with the capacity to simultaneously detect a wide range of microbes.

The first significant effort to develop a microarray for broad-range pathogen identification focused on detecting all known respiratory viruses (32). This microarray contained ∼1,600 oligonucleotides representing ∼140 viruses from all virus families with members known to cause respiratory disease. A more comprehensive array, the ViroChip, was designed using 70-mer oligonucleotides derived from every fully sequenced viral genome in GenBank at the time (33). The oligonucleotide probes selected were highly conserved 70-mers from each viral taxon, enabling the detection of known and unknown family members through cross-hybridization. Following the random amplification of extracted nucleic acids, samples are labeled with fluorescent dyes (typically Cy3 or Cy5). The resulting hybridization patterns on the microarray can be interpreted to infer the presence or absence of known or novel viruses. A variety of approaches for microarray data analysis, including visual inspection, hierarchical clustering (34), and several customized bioinformatics programs, such as GreeneLAMP (35), E-Predict (36), DetectiV (37), PhyloDetect (38), and VIPR (39) and VIPR HMM (40), have been developed. The ViroChip was successfully utilized to identify a novel coronavirus as the causative agent for the severe acute respiratory syndrome outbreak in 2003 (33, 41, 42). Other broadrange DNA microarrays, in addition to the ViroChip, have been described. GreeneChip, a pan-microbial microarray that contains probes for viruses, bacteria, and parasites (35), has been successfully used in a variety of diagnostic scenarios to detect known or unexpected viruses (43–47). Lawrence Livermore Microbial Detection Array is a high-density microarray that contains target probes against every bacterium and virus whose full genome sequence was available at the time of development (48, 49). This array, in combination with multiple displacement amplification-based whole-genome amplification via isothermal amplification by the φ29 polymerase, was successfully used to detect the presence of many DNA and RNA viruses in various types of clinical specimens (50). PathChip is another pathogen microarray option that utilizes random-tagged PCR prior to hybridization, and has been used to identify viral pathogens in nasal wash specimens collected from pediatric patients with respiratory tract infections (51). DNA microarrays can also be used for subtyping of viruses such as influenza A (52). On the bacterial front, microarrays with probes targeting detection of 16S rRNA gene sequences have been developed to define the bacterial diversity present in a given sample. The PhyloChip, a high-density 16S rRNA gene microarray containing ∼300,000 probes, was used to examine microbial diversity in three environmental samples, including urban aerosol, subsurface soil, and subsurface water (53). This study demonstrated that this DNA microarray could reveal a broad range of microbial diversity. A second microarray containing 10,500 probes, including 9,000 taxonomically specific probes targeting the 16S rRNA gene, was used to perform a systematic and quantitative study of bacterial colonization in the infant gastrointestinal tract (54). These microarray results compared favorably with data generated by sequencing-based techniques, demonstrating the utility of microarrays. Microarray-based assays have been successfully utilized to identify bacterial pathogens in several types of infections, such as bacteremia (55–57), genitourinary infections (58, 59), and infectious diarrhea (60). Fungal identification strategies by microarrays have utilized targets such as the 28S rRNA gene and internal transcribed spacer regions within rRNA gene clusters. Microarray technology has been used to identify common fungal

242 n DIAGNOSTIC STRATEGIES AND GENERAL TOPICS

pathogens in humans, including microbes in the genera Candida, Aspergillus, and Fusarium (61, 62). DNA microarrays aimed at microbial diagnostics have benefited immensely from the genomics era, which has spawned a wealth of microbial sequences available in public DNA databases. As demonstrated by successful examples using the ViroChip, microarrays can be a robust platform for virus and pathogen discovery. However, a clear limitation of all hybridization-based strategies for pathogen discovery, including DNA microarrays, is that sequences must be known and well characterized. DNA sequencing strategies, by contrast, offer a relatively openended strategy for pathogen discovery that does not rely on the development of specific probes in advance. The data generated by DNA sequencing platforms can be used as virtual “probes” with DNA databases for identification of unexpected pathogens.

Sequencing-Based Approaches Pathogen discovery and microbial sequencing techniques merged with the discovery of Human metapneumovirus (HMPV) in 2001, which combined classic viral culture with a molecular strategy, termed random arbitrarily primed PCR (63). Efforts to culture respiratory secretions from children suffering respiratory tract infections led to the identification of a putative unidentified virus that could be passaged in several mammalian cell lines. Arbitrary primers were used to generate PCR amplicons, and those amplicons that were uniquely present in infected cells were selectively sequenced by Sanger sequencing methods. Multiple fragments having limited sequence identity to avian pneumoviruses were detected, indicating that a novel virus, now known as HMPV, was present in the infected cells. Seroprevalence studies indicated that by the age of 5 to 10 years, most individuals were antibody positive, suggesting that this virus is a common infection acquired in childhood. HMPV can cause severe respiratory infections similar in presentation and case severity to respiratory syncytial virus, including pneumonia and bronchiolitis, and is responsible for 5 to 10% of hospitalizations of patients with respiratory tract infections (64). In the same year, a candidate independent sequencing strategy for identification of novel viruses, termed DNaseSISPA, was described (65), and it was the methodological basis for discovering novel viruses for nearly a decade. The experimental strategy relied on sequence-independent single primer amplification (SISPA), wherein an adaptor containing a primer-binding sequence is ligated to both ends of a cDNA fragment and a single primer is then used for PCR. To enrich specifically for viral nucleic acids present in virions, the clinical specimen is first subjected to ultracentrifugation to collect the virions, which are subsequently treated with DNase to degrade any cellular nucleic acids that are not protected within the viral capsids. Following this enrichment, the sample is extracted for DNA or RNA and amplified using SISPA. The enrichment steps in this protocol are necessary to increase the chances of sequencing a virus-derived sequence, given the labor and costs of performing extensive Sanger sequencing on the unenriched sample. In this proof-of-concept study, two novel bovine parvoviruses were identified (65). DNase-SISPA and variations of this method were successfully applied to discover many potentially pathogenic viruses with low sequence homology to known viruses during the next several years (Table 1). The identification of these novel viruses highlights the utility of DNA sequencing for viral detection.

Advances in virus discovery continued with the combination of DNase-SISPA-based methods and high-throughput Sanger sequencing. Human bocavirus (HBoV) was discovered in 2005 (66) from respiratory secretions pooled from multiple patients with an unexplained respiratory illness. Amplicons of 600 to 1,500 bp were cloned and sequenced using high-throughput Sanger sequencing with one 384well plate. Phylogenetic analysis demonstrated that this novel genome is a previously uncharacterized species of the genus Bocavirus, HBoV. Subsequent studies of this virus have demonstrated that HBoV is frequently detected in children with respiratory tract infections, children with asthma exacerbations, and children with acute gastroenteritis. Seroepidemiology studies have confirmed infection in a Japanese cohort, with 71.1% overall prevalence with exposure by age 6 (67), while a second study in Sweden reported a lower rate, 33%, in a cohort of children with acute wheezing (68). The application of the same method with multiple specimen types resulted in the discovery of several novel viruses that have yet to be confirmed as pathogens (see Table 1). These discoveries demonstrate that sequenceindependent amplification followed by limited Sanger capillary sequencing (typically ≤384 clones) is a robust method for identification of novel viruses present in clinical specimens. The advent of next-generation sequencing (NGS) technology increased the sequencing depth of specimens, allowing for detection of microbes present at lower titers as well as facilitating the generation of complete genomes of novel microbes. The 2008 discovery of Merkel cell polyomavirus (MCPyV) was the first study describing the identification of a novel virus using NGS on the Roche 454 FLX platform (69). In this instance, cDNA libraries made from Merkel cell carcinoma tumors were sequenced, and from the ∼382,000 high-quality sequence reads generated, one fragment had detectable sequence similarity to a known polyomavirus. Further analysis demonstrated that a highly divergent polyomavirus genome, that of MCPyV, was present in the majority of the Merkel cell carcinoma tumors examined. Subsequent studies have corroborated this finding, and mapping of integration sites demonstrated that the virus was clonally integrated in the respective tumors. Given the very low abundance of MCPyV mRNA sequences in these samples, the detection of viral transcripts would not have been possible without the use of the next-generation platform, which enabled deep sequencing of the specimens in a cost-effective manner. For other novel tumor-associated viruses that have been identified using NGS technology, see Table 2. NGS by 454 FLX has played a pivotal role in defining the etiology of several unexplained infectious illnesses. The first was a mysterious case cluster of five patients with undiagnosed hemorrhagic fever (70). RNAs from two postmortem liver biopsy samples and one serum sample were randomly amplified and sequenced, and the analysis of ∼300,000 sequences yielded nine fragments with limited sequence similarity to viruses in the genus Arenavirus. Phylogenetic analysis of the novel Lujo virus demonstrated that it branched from the Old World arenavirus complex and had the greatest similarity to Mobala virus, Lassa fever virus, and Tamiami virus, with 67 to 74% amino acid identity in the nucleoprotein. Further examination of the receptorbinding portion of G1 demonstrated that Lujo virus is equally distant from the Old World and New World arenaviruses. Other elusive etiologic viral agents of unexplained gastroenteritis have recently been identified using 454 FLX, and they are summarized in Table 2.

16. Microbial Genomics and Pathogen Discovery n TABLE 1

243

Novel viruses discovered by Sanger sequencing methods Virus

Specimen type

Discovery method

Human metapneumovirus

Respiratory secretions

Bovine parvoviruses

Bovine serum

Human coronavirus NL63

Respiratory samples

Parvovirus 4

Plasma

Anellovirus SA1 Anellovirus SA2 Anellovirus Anellovirus Human bocavirus

Plasma Plasma Healthy donor blood Healthy donor blood Respiratory secretions

DNase-SISPA DNase-SISPA DNase-SISPA DNase-SISPA DNase-SISPA + highthroughput Sanger sequencing

KI polyomavirus

Respiratory secretions

WU polyomavirus

Respiratory secretions

Saffold virus

Stool

Astrovirus MLB1

Stool

Human cosavirus A1

Stool

Human bocavirus 2

Stool

Human bocavirus 3

Stool

Bat polyomaviruses

Bat blood, major organs, and rectal and oral swabs

DNase-SISPA + highthroughput Sanger sequencing DNase-SISPA + highthroughput Sanger sequencing DNase-SISPA + highthroughput Sanger sequencing DNase-SISPA + highthroughput Sanger sequencing DNase-SISPA + highthroughput Sanger sequencing DNase-SISPA + highthroughput Sanger sequencing DNase-SISPA + highthroughput Sanger sequencing Seminested PCR + Sanger sequencing

Viral culture, random arbitrarily primed PCR, and Sanger sequencing DNase-SISPA

Virus discovery cDNAamplified fragment length polymorphism DNase-SISPA

Sequencing-based discovery of novel bacterial pathogens can be performed by sequencing specific gene targets or using a combination of whole-genome shotgun sequencing (WGS) and a range of data analysis tools. These tools may be applied to the assembly and analysis of bacterial genomes from either pure culture or patient specimens. A workflow for implementing bacterial WGS-based molecular diagnostics is depicted in Fig. 2. In 2013, a novel species of Bartonella was identified from a patient with chronic bartonellosis by combining culture methods, PCR amplification, and Sanger sequencing of three gene fragments important for speciation. Blazes et al. targeted the rrs, gltA, and rpoB genes, and based on sequence analysis using BLAST, ClustalW, and MEGA5 software, they showed that the sequence similarities to other known Bartonella spp. were below the standard similarity ranges for Bartonella, and identified this novel isolate as “Candidatus Bartonella ancashi” (71). In the same year,

Disease

Reference(s)

Severe respiratory infections

63

Contaminants of commercial bovine serum Croup

65

111

Viral sepsis of immunocompromised patients Unknown Unknown Unknown Unknown Respiratory tract infections, asthma exacerbation, and acute gastroenteritis Unknown

112

Unknown

115

Unknown

13, 116

Unknown

117

112 112 113 113 66

114

Nonpolio acute flaccid paralysis

118–120

Nonpolio acute flaccid paralysis and acute gastroenteritis Acute gastroenteritis

121, 122

Unknown

121

96

Bhatt et al. (19) applied WGS to DNA isolated from formalin-fixed, paraffin-embedded biopsy specimens from patients with cord colitis in an effort to identify a novel etiologic agent for the disease. Previous theories for the etiology of cord colitis assumed it was a manifestation of graft-versushost disease as opposed to a distinct clinical syndrome. Their efforts resulted in the identification of a novel bacterial draft genome, Bradyrhizobium enterica, which was absent from control specimens and associated with all cord colitis specimens they analyzed (Fig. 3). NGS has also been applied to areas of molecular epidemiology and pathogen emergence. In 2011, Wright et al. (72) used NGS to quickly complete the whole-genome characterization of an anthrax-like agent within days of its recovery from antemortem cultures. The etiologic agent was determined to be Bacillus cereus, not B. anthracis. Rapid genome sequencing and analysis of the causative agent of this fatal,

244 n TABLE 2

DIAGNOSTIC STRATEGIES AND GENERAL TOPICS Novel viruses discovered by NGS methods Virus

Specimen type

Merkel cell polyomavirus Epstein-Barr virus type 1

CNSa lymphoma viruses (Epstein-Barr virus, cytomegalovirus, JC virus, and HIV) Lujo virus Human klassevirus 1 Astrovirus VA1 Middle East respiratory syndrome coronavirus Mamastrovirus, Bocavirus, Circovirus, Iflavirus, and Orthohepadnavirus Miniopterus schreibersii papillomavirus

Merkel cell carcinoma tumors Nasopharyngeal carcinoma tumor biopsy RNA from primary CNS lymphomas

RNA from postmortem liver biopsy Pediatric stool Stool Tracheal aspirates and nasopharyngeal or throat swabs Bat thoracic and abdominal organ specimens Bat rectal and oral swabs

Discovery method

Disease

Reference(s)

Roche 454 FLX

Merkel cell carcinoma

69

Illumina Genome Analyzer IIx

Nasopharyngeal carcinoma

123

Next-generation transcriptome sequencing

CNS lymphomas

124

Next-generation transcriptome sequencing Roche 454 FLX GS-FLX Titanium Illumina

Hemorrhagic fever

70

DNase-SISPA + Solexa sequencing

Unknown

95

454 sequencing

Unknown

97

Diarrhea Acute gastroenteritis Middle East respiratory syndrome

a

CNS, central nervous system.

FIGURE 2 A proposed workflow for bacterial, WGS-based molecular diagnostics. This workflow uses benchtop sequencing for data generation and implements cloud storage for central data storage and remote data processing. Double-headed arrows indicate constant updating of the central data repository and reference database. Reprinted from reference 128 with permission of Nature Publishing Group. doi:10.1128/9781555817381.ch16.f2

125, 126 127 18

16. Microbial Genomics and Pathogen Discovery n

245

FIGURE 3 Phylogenetic analysis and genome assembly of Bradyrhizobium enterica, a novel organism associated with cord colitis. Assembly of the B. enterica genome included unmappable reads generated from WGS of samples from patients with cord colitis. (A) Multisequence alignment of 400 core protein-coding genes was used to generate a rooted phylogenetic tree showing the predicted evolutionary relationships between B. enterica and related species. (B) Genomic comparison of B. enterica to Bradyrhizobium japonicum. Dark blue lines on the inner circle depict genes present in the genome of B. enterica that are absent in the genome of B. japonicum. The outer circle depicts amino acid sequence identity of each B. enterica predicted protein to its closest homologue in B. japonicum. The middle circle is a representation of the contig assembly, with the borders of each contig (dark green) outlined in light green. Reprinted from reference 19 with permission of Massachusetts Medical Society. doi:10.1128/9781555817381.ch16.f3

anthrax-like pneumonia effectively determined the etiology of the infection. WGS was first applied as a genome-based molecular epidemiological tool in 2013. Roetzer et al. (73) assessed WGS-based genotyping as a means to track the spread of M. tuberculosis in a metropolitan area and follow its decade-long clonal expansion to determine the shortterm evolution of the M. tuberculosis genome. The results showed that genotyping of an M. tuberculosis outbreak using WGS methods was more robust than conventional methods like IS6110 restriction fragment length polymorphism, spoligotyping, and mycobacterial interspersed repetitive-unit– variable-number tandem-repeat typing, which examine 80%, are considered three distinct species because they differ in many phenotypic and chemotaxonomic aspects (16). It is essential that the boundaries of species demarcation be flexible in order to achieve a classification scheme that facilitates identification. The application of numerous other types of analyses of genotypic, chemotaxonomic, and phenotypic characteristics of bacteria to the delineation of bacteria at various hierarchical levels represents the third component of polyphasic taxonomy (9). The goal is to collect as much information as possible and to evaluate all results in relation to each other in order to draw useful conclusions. An additional advantage is that once the taxonomic resolution of these approaches has been established for a particular group of bacteria through the analysis of taxonomically well-characterized strains, they may be used as alternative tools to identify new isolates at different taxonomic levels. It should be noted that the resolution of these alternative methods is often group dependent. For instance, cellular fatty acid analysis is useful for the accurate identification of strains of many bacterial species to the species level. In certain bacterial groups, however, the cellular fatty acid profile may be indicative of the genus or a group of phylogenetically related genera but not of a particular species within one of these genera. The contours of a polyphasic bacterial species are obviously less clear than the ones defined by Wayne et al. (12), and this lack of a rigid definition has been contested, as it allows too many interpretations (10, 17, 18). Polyphasic classification is empirical and contains elements from both phenetic and phylogenetic classifications. There are no strict rules or guidelines, and the approach integrates any significant information on the organisms, resulting in a consensus type of classification. In this respect, its main weakness is indeed that it relies on common sense to draw its conclusions. The bacterial species appears as a group of isolates in which a steady generation of genetic diversity resulted in clones characterized by a certain degree of phenotypic consistency, by a significant degree of DNA-DNA hybridization, and by a high level of 16S rRNA sequence similarity.

17. Taxonomy and Classification of Bacteria n

The species is the most important and, at the same time, the central element of bacterial taxonomy. There are at present no rules for the delineation of higher hierarchical ranks such as genus, family, and order. Although there is an expectation that at the generic level taxa should be supported by phenotypic descriptions (5, 10, 13), in practice, higher ranks are mostly delineated on the basis of 16S rRNA sequence comparison and stability analyses of the clusters that are obtained (19, 20). The latter has weakened the emphasis on phenotypic descriptions of taxa. Nevertheless, analyses of conserved signature indels and conserved signature proteins demonstrate that groups of prokaryotes ranging from phylum to genus levels can be recognized and support a tree-like vertical inheritance of the genes containing these molecular signatures that is consistent with phylogenetic trees (21).

Multilocus Sequence Analysis: A Short-Lived Alternative to DNA-DNA Hybridization Experiments? In 2002, a new ad hoc committee for the reevaluation of the species definition in bacteriology made various recommendations in light of developments in methodologies available to systematists (22). One of the particularly interesting developments was multilocus sequence analysis (MLSA). In contrast with multilocus sequence typing (MLST), a specific tool designed for molecular epidemiology and for defining strains within named species, whereby similarities and differences are usually measured as differences in allelic profiles, MLSA employs phylogenetic procedures based on the nucleotide sequences of the alleles instead to reveal similarities between strains representing different species and genera (6, 23). Many examples of such studies have been published, and in general the clusters delineated correlated well with species demarcated by DNA-DNA hybridization experiments (6). It is, for instance, noteworthy that the DNA-DNA hybridization results that demonstrated that Yersinia pestis and Yersinia pseudotuberculosis represented a single species were mirrored in the MLSA tree, where Y. pestis clusters among Y. pseudotuberculosis strains; the same observation was made for Burkholderia mallei and Burkholderia pseudomallei (24, 25). There is no universal MLSA cutoff or descriptor of clusters that characterizes species, nor are ecological features consistently available to distinguish natural clusters that could be used to define species. Therefore, MLSA thresholds for species delineation must be validated through the analysis of taxonomically well-characterized reference strains (13) before this method can replace DNA-DNA hybridization experiments in the frame of polyphasic taxonomic studies (see, e.g., reference 26). Especially for depicting relationships within and between closely related species, this approach has a resolution superior to the traditional 16S rRNA gene sequence analysis. The deduced phylogenetic trees not only provide a phylogenetic backbone but also reveal intraspecies relationships at a level where comparative 16S rRNA sequence analysis is no longer discriminatory. The number and lengths of gene fragments to be used in MLSA studies have not been systematically studied, although typically six to eight genes are analyzed (27). Housekeeping genes are preferentially used because they evolve relatively slowly and most of the variation that accumulates in these genes is considered selectively neutral. Sequence diversity, however, often precludes the development of primer sets that can be used for studying multiple genera

257

or even species. Although very appealing for its resolution, portability, and throughput capacity, MLSA thus suffers from the difficulty of developing widely applicable schemes. As the costs of whole-genome sequencing continue to decline, it is conceivable that extracting MLSA sequences (or MLST alleles) from draft genome sequences will become more straightforward and affordable than performing traditional MLST or MLSA analyses (28).

Toward a Genomic Threshold for Species Definition For several decades, bacterial taxonomists have considered whole-genome information the standard for determining taxonomy. The number of whole-genome sequences is increasing rapidly and allows assessment of genome-level variation within and between species. It has become clear that in addition to nucleotide substitutions, other genetic forces such as gene loss, gene duplication, chromosomal rearrangements, but also LGT shape the genome and that considerable fractions of the genome of any particular strain may be unique to that strain (29). One of the more significant recent discoveries in bacterial genomics is that bacterial species appear to comprise a set of core and accessory genes, with only the former present in all isolates of that species and with the sum of the two components forming the species pan-genome (30). The origin, composition, and size of bacterial pan-genomes and whether they are finite or infinite have been the source of debate (31). In particular, the extent to which LGT occurs and its consequences for bacterial evolution, a biological species concept of bacteria, and eventually a bacterial species definition have been debated vigorously. Although recognized decades ago, LGT was long viewed as a minor phenomenon that did not jeopardize the general idea of vertical evolution as depicted in phylogenetic trees based on rRNA or other conserved genes. An overwhelming disruptive influence of LGT also did not correspond with daily diagnostic practices during which numerous bacterial isolates can be recognized as belonging to long-established species. However, MLSA and especially phylogenomics studies have shown that numerous genes have different evolutionary histories, and the ribosomal tree of life has been referred to as “the tree of one percent” (of all genes in microbial genomes) (32, 33). Microbial evolution seemed better modeled in the form of a dynamic network of evolution in which the nodes were bacterial and archaeal genomes and the edges were the fluxes of genetic information between the genomes (33). Yet there is now compelling evidence that a considerable set of genes share a highly significant phylogenetic signal that can be used to reconstruct bacterial phylogeny (34). The traditional tree of life can be replaced with a common and coherent phylogenetic trend of many genes, called the phylogenetic forest or the statistical tree of life (33, 34). Several studies of complete genomes have suggested universal sets of protein-coding genes that may be useful for a phylogenomic species delineation in microbial taxonomy, and genes that encode components of the translation system in particular show substantial congruency between each other and with the standard rRNA tree (35–39). It also became apparent that not only does the frequency of LGT vary dramatically among bacteria (an observation that was apparent more than a decade ago through numerous DNA fingerprinting studies) but that LGT should be considered a cohesive force in evolution rather than a disruptive force leading to a genetic melting pot (18, 40, 41), as the requirement for physical proximity, the homology dependence for

258 n

BACTERIOLOGY

successful recombination, and the fitness reduction in the recipient organism all favor recombination between closely related bacteria (40). Finally, the differential existence of an accessory genome is clearly compatible with LGTdriven environmental adaptation. In a study of the evolutionary origin of the genomic repertoires in Gammaproteobacteria, Lerat et al. (42) concluded that gene acquisition was very common based on the large number of genomeor clade-restricted gene families. However, beyond their initial acquisitions, few gene histories conflicted with the organismal tree. The picture emerged that bacterial lineages are constantly subjected to the input of new genes from a large available pool, which may be of bacteriophage origin, and that resident genes are continually lost. The diversity of gene families unique to single genomes indicates that the pool of available genes is very large, allowing the rate of gene acquisition to be both high for a genome and very low for a particular gene. A taxonomically particularly interesting study was published by Lefebure et al. (43), who analyzed 96 genome sequences derived from two closely related sympatric sister species of pathogenic bacteria, Campylobacter coli and Campylobacter jejuni. The results showed that both species had finite pangenomes and that there are unique and cohesive features to each of their genomes defining their genomic identity. The two species have a similar pan-genome size; however, C. coli has acquired a larger core genome and each species has evolved a number of species-specific core genes, possibly reflecting different adaptive strategies, in spite of their occurrence in the same niche (the gastrointestinal tract of several hosts). Genome-wide assessment of the level of LGT within and between the two sister species, as well as within the core and noncore genes, demonstrated a resistance to interspecies recombination in the core genome of the two species and therefore provided persuasive support for the core genome hypothesis for bacterial species. How can all of this information be used in a new species definition? Studies by Konstantinidis and Tiedje (29) and Goris et al. (44) revealed the average nucleotide identity (ANI) of the shared genes between two strains to be a robust means to compare genetic relatedness. ANI values of ∼95% corresponded to the traditional 70% DNA-DNA hybridization standard of the current species definition, and several recently published descriptions of novel bacterial species have incorporated the ANI analysis of draft wholegenome sequences (a minimum of 20% of the randomly sequenced complete genome must be available) to replace DNA-DNA hybridization experiments (5, 45). At the 95% ANI cutoff, current species include only moderately homogeneous strains, apparently as a result of the strains having evolved in different ecological settings. A large fraction of the differences in gene content within species was associated with bacteriophage and transposase elements, revealing an important role of these elements during bacterial speciation. These findings were consistent with a definition for species that would include a more homogeneous set of strains than that provided by the current definition and one that considers the ecology of the strains in addition to their evolutionary distance (46). Goris et al. (44) also demonstrated that the 70% DNA-DNA reassociation threshold corresponded with 69%-conserved DNA, or, when the analysis was restricted to the protein-coding portion of the genome, 85%-conserved genes. ANI values detect the DNA conservation of the core genome, whereas conserved DNA calculates the proportion of DNA shared by two genomes. Both estimates of intraspe-

cies similarity therefore do not necessarily correlate, and for that reason, Deloger et al. (62) introduced the maximal unique match (MUM) index values, which take into account both criteria of diversity. MUM index values represent a calculation for genomic distances that is based on the number of maximal unique and exact matches of a given minimal length shared by the two genomes being compared. MUM index values correlate better with the ANI than with the DNA content and group strains in a way that is congruent with MLSA trees.

Major Groups of Bacteria The tree of life, based on comparative small-subunit rRNA studies, comprises three lines of descent that are nowadays referred to as the domains Bacteria, Archaea, and Eucarya (8). There are currently 34 formally named phyla in the Bacteria, which are further subdivided into numerous taxa (47; http://www.ncbi.nlm.nih.gov/Taxonomy/tax.html/). However, a large number of uncultured bacteria represent additional phylogenetic lineages awaiting formal classification, and alternative phylogenetic classification systems (and thus alternative nomenclatures) have been proposed in which, for instance, more-comprehensive data sets, including multiple molecular sequences and cell envelope and ultrastructural characteristics, rather than reliance solely on rRNA sequence similarity, define the units of classification (48). The present division into phyla is largely based on the classification proposed in the second edition of Bergey’s Manual of Systematic Bacteriology (20). However, Bergey’s Manual of Systematic Bacteriology is not an official document endorsed by the International Committee on Systematics of Prokaryotes, and the nomenclature and classification schemes presented are no more than recommendations by the editors and authors of that manual. There is no official or valid publication of classification schemes in bacterial taxonomy, and the International Committee governs only the rules of nomenclature of prokaryotes and matters relating to the Bacteriological Code (47). Recently, Gribaldo and Brochier-Armanet (49) called for a general discussion on the higher taxa of prokaryotes, to lay down common rules for the establishment of names for such taxa and to broaden the scope of the International Code of Nomenclature of Prokaryotes to include the higher taxa. In addition to 16S rRNA-based distinctiveness, the use of biological signatures, such as the occurrence of unique features in terms of key cellular processes identified by distinctive gene distributions and the availability of at least one cultivated representative and a complete genome sequence, was recommended to define phyla (49). In the traditional (rRNA gene sequence-based) view of higher-order taxonomy and phylogeny, three phyla, the Proteobacteria, the Firmicutes (Gram-positive organisms with low G+C contents, including Bacillus, Clostridium, Staphylococcus, Mycoplasma, and the classical lactic acid bacteria, such as Enterococcus, Streptococcus, and Lactobacillus), and the Actinobacteria (Gram-positive organisms with high G+C contents, including Bifidobacterium, Mycobacterium, and Corynebacterium), comprise the large majority of clinically relevant species. The Bacteroidetes (Bacteroides, flavobacteria, and sphingobacteria), the Spirochaetes (spirochetes and leptospiras), and the Chlamydiae (chlamydias) represent some of the other phyla. A detailed overview is given by Garrity and Holt (20) in their introductory chapter to the second edition of Bergey’s Manual of Systematic Bacteriology. That edition is structured in an order based on the topology of the 16S rRNA phylogenetic tree.

17. Taxonomy and Classification of Bacteria n 259

The largest phylum by far is the Proteobacteria, which contains five main clusters (classes) of genera that are referred to with the Greek letters alpha, beta, gamma, delta, and epsilon. More recently, two additional classes within the Proteobacteria have been reported, the “Zetaproteobacteria” (a name that has not been validly published and therefore is formally written between quotation marks) and the Acidothiobacillia (50, 51). The Proteobacteria comprise the majority of the known Gram-negative bacteria of medical, industrial, and agricultural significance. This phylum includes Brucella, Ehrlichia, and Rickettsia (Alphaproteobacteria); Burkholderia, Bordetella, and Neisseria (Betaproteobacteria); Aeromonas, Legionella, Vibrio, and the family Enterobacteriaceae (Gammaproteobacteria); and Campylobacter and Helicobacter (Epsilonproteobacteria). The Deltaproteobacteria, “Zetaproteobacteria,” and Acidothiobacillia comprise a variety of mainly environmental bacteria that have little clinical relevance.

Uncultured Bacteria The classification and nomenclature of uncultured bacteria that are only minimally characterized by morphological characteristics or by differences in molecular sequence are outstanding challenges in bacterial classification (52). A category that formally classifies incompletely described prokaryotes has been recognized (53). “Candidatus” is considered a taxonomic status for uncultured candidate species for which relatedness has been determined (for instance, for which phylogenetic relatedness has been determined by amplification and sequence analysis of prokaryotic RNA genes by use of universal prokaryotic primers) and whose authenticity has been verified by in situ probing or a similar technique for cell identification. In addition, it is also mandatory that information concerning phenotypic, metabolic, or physiological features be made available. The latter data may serve as a starting point for further investigation and eventual description and naming. With the advent of genomics, these original concerns may appear trivial now that we have the technical means to study large genomic fragments of uncultured microbes by shotgun cloning and sequencing of bulk DNA extracted from mixed communities (54, 55) or by DNA amplification from single cells (56), enabling sequencing from uncultured microorganisms from the environment (57). In particular, the metagenomics field has opened a fascinating new window for studying the uncultured microbial diversity in a range of ecosystems, including those of the human body (see chapter 15). Exploring the diversity of such ecosystems by comparative rRNA sequence analysis is based on identification of phylotypes, which are defined as groups of 16S rRNA gene sequences with a certain level of similarity. The cutoff values of rRNA sequence similarity that are used for phylotype definition are not consistent. In studies of the diversity of microbes in the human gastrointestinal tract, these values vary between 97 and 99% (58). The higher the cutoff value, the higher is the number of distinct phylotypes and thus estimated species richness. Regardless of the cutoff value used, the resulting diversity estimates should be considered rough indicators of the microbial diversity present, as bacteria with rRNA sequences that are 99% similar may still encompass multiple species and considerable ecological and genomic heterogeneity (6).

(species, genus, or higher taxonomic rank) and designated accordingly. It relies on a comparison of the characters of an unknown with those of established units in order to name it appropriately. This implies that identification depends on adequate characterization. As part of identification strategies, dichotomous keys based on morphological and biochemical characteristics have only partly been replaced by other methods. Taxonomic studies provide an impressive array of alternative techniques derived from analytical biochemistry and molecular biology for examination of numerous cellular compounds (11, 13). Various identification approaches are discussed in chapters 4 through 6 of this Manual. Each of these techniques is useful for characterization and hence identification of bacteria. Databases of many types of biological data, including rRNA and other gene sequences, wholecell fatty acid components, and matrix-assisted laser desorption ionization–time-of-flight (MALDI-TOF) mass spectra, or miniaturized series of phenotypic characteristics may allow identification of many isolates. Yet the success of these databases also depends on the completeness of the databases, the exactness of the methods, and how carefully the individual entries have been delineated.

CLASSIFICATION AND IDENTIFICATION METHODS In principle, all genotypic, phenotypic, and phylogenetic information can be used to classify bacteria. Genotypic information is derived from the nucleic acids present in the cell, whereas phenotypic information is derived from proteins and their functions, different chemotaxonomic markers, and a wide range of other expressed features. In the present polyphasic-species definition, a minimal taxonomic study consists of sequence analysis to determine the phylogenetic position of the unknown species (the 16S rRNA gene is most commonly used), DNA-DNA hybridizations to determine its precise level of relatedness toward its nearest phylogenetic neighbors, and biochemical characterization to distinguish the new taxon from the established ones. Typical for the process of polyphasic taxonomy is that information from other approaches is used to classify bacteria at different taxonomic levels. When working one’s way through lists of methods, it is of primary interest to understand at which level these methods carry information and to realize their technical complexity, i.e., the amount of time and work required to analyze a certain number of isolates. The validation of a new classification or identification tool involves a determination of its taxonomic resolution by means of well-characterized reference strains. As a proof of concept, such validation studies mostly start with the analysis of type strains only, but because of the genotypic variability of bacterial species (both in gene content and in gene sequences), the true value of new classification or identification tools can appropriately be assessed only through the analysis of multiple well-characterized reference strains and subsequent validation using new isolates. The list of methods given below is not meant to be complete or to describe all of their aspects. It comprises the major categories of taxonomic techniques required to classify and identify bacteria and focuses on novel developments.

DNA-DNA Hybridization Studies

IDENTIFICATION Identification is part of taxonomy. It is the process whereby an organism is recognized as belonging to a known taxon

Although contested for its technical difficulties and for the inability to build cumulative databases, at present, DNA-DNA hybridization is still acknowledged as the reference method to establish relationships within and

260 n

BACTERIOLOGY

between species. Different DNA-DNA hybridization procedures have been described, both traditional and miniaturized versions (9, 59–61). In DNA-DNA hybridization studies, it is commonly unclear if hybridizations were performed under optimal, stringent, or suboptimal conditions. The stringency of the reaction is determined by the salt and formamide concentrations, by the temperature, and by the molar percentages of G+C of the DNAs used. DNA-DNA hybridizations performed under standard conditions are not necessarily optimal or stringent for all bacterial DNAs. As a standard, optimal conditions for hybridizations should be preferred because the optimal temperature curve for hybridization is rather broad (∼5°C) (9).

Whole-Genome Sequence-Based Methods The percentage of conserved DNA (pcDNA) method introduced by Goris et al. (44) and the MUM index of Deloger et al. (62) can be regarded as in silico imitations of DNADNA hybridization. Both similarity measures are based on finding subsequences that are conserved in the genomes under comparison. However, pcDNA traces inexact matches using a BLAST approach (63), whereas the MUM index uses the MUMmer software (62) to find all maximal unique exact matches (MUMs) shared by the genomes. MUMmer applies suffix trees as an index structure to speed up the search process. As a result, computing the MUM index is ∼100 times faster than the pcDNA method (64). Both methods require complete or nearly complete genome sequences, as they are highly sensitive to large sequence stretches missing in draft genomes. The ANI method (5, 44, 65) and core gene identity (CGI) method (66) are both natural extensions to MLSA in case complete or draft genome sequences are available. Whereas MLSA computes similarity values as the average similarity among a limited set of orthologous protein-coding genes shared among all organisms studied, ANI and CGI maximize the information content by taking all orthologous protein-coding genes of the genomes into consideration. The main difference between the approaches is that ANI computes a pairwise genome similarity based on all orthologous genes shared between two genomes, while CGI restricts the computation of pairwise genome similarity to orthologous genes shared by all genomes under study. The present state of the art demonstrates that both pcDNA and MUM index parameters are equivalent (because they have the same taxonomic resolution) but superior (because they are sequence based and thus cumulative) approaches to the traditional DNA-DNA hybridizations for the species-level classification of bacteria. Similarly, the ANI and CGI parameters are equivalent but superior tools for studying their phylogeny. The latter techniques are superior compared with traditional 16S rRNA-based approaches because they are based on a much larger part of the genome and because they have a better resolution for discriminating closely related bacteria. Each of these methods is starting to be used in taxonomic studies of different bacteria (45, 64, 66, 67), and it can be anticipated that the practice of DNA-DNA hybridization experiments as the cornerstone for species delineation in taxonomic studies of bacteria will be abandoned fairly soon.

rRNA Studies rRNA is the best single target for studying phylogenetic relationships because it is present in all bacteria, it is functionally constant, and it is composed of highly conserved as well as more variable domains (8). The compo-

nents of the ribosome (rRNA and ribosomal proteins) have been the subjects of phylogenetic studies for several decades, and direct sequencing of partial or nearly entire 16S or 23S rRNA molecules has become common practice. The larger the conserved elements examined, the more information they bear and the more reliable the conclusions become. International databases comprising published and unpublished partial or complete sequences have been constructed (68) but have also accumulated poorquality sequences and sequences that are not accurately or even not correctly labeled. For these and other reasons, several initiatives for providing the scientific community with curated 16S rRNA databases have been undertaken (e.g., see references 69–71). The “All-Species Living Tree” project (69; http://www.arb-silva.de/projects/living-tree/) aims to reconstruct a single 16S rRNA tree harboring all sequenced type strains of the hitherto classified species of Archaea and Bacteria. Sequences are selected manually due to a high error rate in the names and information fields provided for the publicly deposited entries. A most useful tool to track the identity of strains for which sequences are deposited is the StrainInfo bioportal (72; http://www.straininfo.net/), which brings together the biological material kept at multiple biological resource centers into a single portal interface, with direct pointers to the relevant information at the culture collections’ websites. This information is automatically linked to related sequences in the public domain and refers to all known scientific publications that deal with the organism. To support the taxonomic depth of the information provided by the StrainInfo bioportal, all taxonomic names appearing in the bioportal are fully integrated with and linked out to key taxonomic information sources. rRNA sequence analysis not only is used to determine relationships among genera, families, and other higher ranks but often replaces DNA-DNA hybridization studies for the delineation of species in taxonomic practice. Such application of rRNA similarity data may not be appropriate, as some genera, for instance, Bacillus, Burkholderia, Corynebacterium, Mycobacterium, and many others, comprise clusters of closely related species that exhibit nearly identical 16S rRNA gene sequences. Stackebrandt and Goebel (73) reported that organisms sharing >98.5% rRNA similarity may or may not belong to a single species and that the resolution of 16S rRNA sequence analysis for determination of the degree of relatedness between closely related organisms is generally low. There is no single threshold value of 16S rRNA similarity for species recognition (73). However, organisms with 60%, no matter which DNADNA hybridization method is used. Subsequent studies revealed that for the majority of organisms, the 97% cutoff value could be raised to 98.7 to 99% (74). Nevertheless, other studies extended the observations on intraspecies 16S rRNA divergence considerably, as differences in 16S rRNA gene sequence of up to 4.5% were reported among strains of several species belonging to the Epsilonproteobacteria (75, 76). In spite of its limitations, rRNA sequence analysis is commonly used for the identification of bacteria (77, 78), and commercial identification systems based on analysis of rRNA gene sequences are available (e.g., MicroSeq 500 16S rDNA Bacterial Sequencing Kit; Perkin-Elmer Applied Biosystems, Foster City, CA). A fraction of the 5′-terminal region of the 16S rRNA gene (positions 60 to 110 of the E. coli numbering system) is one of the most informative or discriminating regions for closely related organisms (19).

17. Taxonomy and Classification of Bacteria n

Comparison of 16S rRNA gene sequences in many bacterial genera will lead to correct identification to the species level, but it is equally true that many taxonomic studies have revealed that comparative rRNA sequence analysis is often not sensitive enough to identify strains to the species level. There is a lack of knowledge not only of the strain-tostrain variation within a species but also of the interoperon variation within a single strain. Therefore, concluding that an unidentified isolate belongs to a particular species because it shares a high percentage of its 16S rRNA gene sequence with particular species or concluding that it represents a novel species because it occupies a unique position in the phylogenetic tree supported by a high bootstrap value or because it shares only 97% of its 16S rRNA sequence with its closest neighbor is premature in the absence of appropriate complementary data. This is even more true for partial sequence data, as partial rRNA gene sequences carry only limited information about the molecule and different parts of the gene may carry information about different taxonomic levels. The interesting taxonomic properties of rRNA or rRNA gene molecules have been exploited in many ways. Although highly conserved, the rRNA genes also consist of variable domains that are particularly useful for diagnostic purposes and for mixed-community analyses (79–82). New technologies, exploiting the universal characteristics of the rRNA genes and their potential for species identification, emerge regularly. A growing number of studies report on the use of pyrosequencing (Biotage, Uppsala, Sweden), which provides rapid, short-read sequencing of 30 bases to classify, identify, and subtype bacteria, yeasts, and fungi (see, e.g., references 83 and 84). Turenne et al. (85) used single-stranded conformation polymorphism analysis of PCR amplicons to distinguish between organisms, and Yang et al. (86) used high-resolution melt analysis to characterize PCR products generated from three hypervariable regions of the 16S rRNA gene of clinically relevant bacterial pathogens and concluded that it allowed highly specific species identification. Still other approaches combine the diagnostic potential of 16S rRNA genes with the speed and discriminatory power of mass spectrometric analyses (see below).

261

genome sequences have suggested universal sets of proteinencoding genes (35–39), and the analysis of multiple protein-encoding genes (not necessarily the most conserved ones) buffers the distorting effects of horizontal gene transfer or recombination events (65).

Other Genotypic Methods for Bacterial Classification A range of different genotypic techniques has been used to characterize bacteria at various taxonomic levels. The molar percentage of guanine plus cytosine (the DNA base ratio or percent G+C value) is one of the classical genotypic characteristics and is part of the standard description of bacterial taxa. Generally, the range in G+C content observed among the strains of a species should not be >3%, and among the species of a genus it should not be >10%. In general, in the bacterial world the G+C content varies between 24 and 76%. A tremendous number of molecular diagnostic methods, most of which are PCR based, have been developed. Most of these generate arrays of DNA fragments that are separated and detected in various ways, and appropriate software has been developed for pattern recognition and analysis and for database construction. One of these DNA fingerprinting methods, amplified fragment length polymorphism analysis (94), is useful for the classification of strains at the species and genus levels. The basic principle of amplified fragment length polymorphism analysis is restriction fragment length polymorphism analysis, modified by using PCR-mediated amplification to select particular DNA fragments from the pool of restriction fragments. This selective amplification process results in an array of about 30 to 40 DNA fragments, some of which are species (or even genus) specific, while others are strain specific (95). PCR-based typing methods that use random or repetitive elements as primers have been applied to strain characterization of a wide variety of bacteria (96–98). Although primarily applied for infraspecies strain comparisons, these techniques are useful in classification as well.

Phenotypic Methods Sequence Analysis of Protein-Encoding Genes As an alternative to 16S rRNA gene sequence analysis, numerous other macromolecules have been examined for their potential as microbiological clocks and their applications as identification tools. Among others, various ribosomal proteins (37, 87, 88), chaperonin (88), RNA polymerases (89), RecA (90), and manganese-dependent superoxide dismutase (91) were shown to be valuable molecular chronometers. These alternative macromolecules should be widely or universally distributed among bacteria, they should not be transmitted horizontally, and their molecular evolution rate should be comparable to or somewhat higher than that of 16S rRNA, which would render them more suitable for differentiation of closely related organisms. Potential pitfalls of overreliance on a single phylogenetic marker are illustrated in the taxonomic studies of species of the Streptococcus bovis group. Streptococcus infantarius subsp. coli was reclassified as the novel species Streptococcus lutetiensis, and another group of streptococci was proposed as the novel species Streptococcus pasteurianus, primarily on the basis of manganese-dependent superoxide dismutase gene sequences (92), while subsequent studies demonstrated that neither S. lutetiensis nor S. pasteurianus represented novel species (93). As mentioned above, studies of complete

Phenotypic methods comprise all those that are not directed toward DNA or RNA and therefore also include the chemical or chemotaxonomic techniques. The classical phenotypic tests traditionally constituted the basis for the formal description of bacterial species, subspecies, genera, and families. While genotypic data are used to allocate taxa to a phylogenetic tree and to draw the major borderlines in classification systems, phenotypic consistency is required to generate useful classification systems and may therefore influence the depth of a hierarchical line (12, 22). The paucity or variability of phenotypic characteristics for certain bacterial groups regularly causes problems in describing or differentiating taxa. For such bacteria, alternative chemotaxonomic or genotypic methods are required to reliably characterize strains. The classical phenotypic characteristics of bacteria comprise morphological, physiological, and biochemical features. Individually, many of these characteristics are poor parameters for genetic relatedness, yet as a whole, they provide descriptive information for the recognition of taxa. The morphology of a bacterium comprises both cellular (shape; the presence of an endospore, flagella, and inclusion bodies; and Gram staining characteristics) and colonial (color, dimensions, and form) characteristics. The

262 n

BACTERIOLOGY

physiological and biochemical features comprise data on growth at different temperatures; growth in the presence of different pH values, salt concentrations, or atmospheric conditions; growth in the presence of various substances, such as antimicrobial agents; and the presence or activities of various enzymes and utilization of compounds, etc. Very often, highly standardized procedures are required to obtain reproducible results within and between laboratories. Phenotypic data were the first to be analyzed by means of computer-assisted numerical comparison. In the 1950s, numerical taxonomy arose in parallel with the development of computers (99) and allowed comparison of large numbers of phenotypic traits for large numbers of strains. Data matrices showing the degree of similarity between each pair of strains and cluster analyses resulting in dendrograms revealed a general picture of the phenotypic consistency of a particular group of strains. Because such large numbers of characteristics reflect a considerable amount of genotypic information, it soon became evident that numerical analysis of large numbers of phenotypic characteristics was indeed taxonomically relevant. In taxonomic practice, phenotypic characterization became compromised and sometimes more of a burden than a useful taxonomic activity. Frequently, phenotypic data are compared with literature data obtained using other conditions or methods. The need for continued phenotypic characterization at every taxonomic level not only to delineate taxa and appreciate their phenotypic coherence but also to evaluate their physiological and ecological functions cannot be denied. A minimal phenotypic description is not only the identity card of a taxon but also a key to its biology. Although accepted as necessary, differential phenotypic characters are often hard to find with a reasonable amount of effort and time.

Chemical Methods The term “chemotaxonomy” refers to the application of analytical methods to the collection of information on various chemical constituents of the cell to classify bacteria. As with the other phenotypic and the genotypic techniques, some of the chemotaxonomic methods have been widely applied to vast numbers of bacteria, whereas others were so specific that their application was restricted to particular taxa. The markers studied include whole-cell protein profiles, isoprenoid quinones, cytochromes, peptidoglycans, polyamines, polar lipids, pigments, particular enzymes, sterols, and hopanoids (11). Very often, analytical difficulties have been the main restrictions to their wide-scale application.

Cell Wall Composition The distinction between Gram-negative and Gram-positive types of bacteria is still one of the characteristics that are first analyzed in order to guide subsequent characterization and identification steps. The determination of the cell wall composition has traditionally been important for Grampositive bacteria. The peptidoglycan type of cell wall of Gram-negative bacteria is rather uniform and provides little information. The cell walls of Gram-positive bacteria, in contrast, contain various peptidoglycan types that may be genus or species specific (100). The most valuable information is derived from the type and composition of the peptide cross-link between adjacent chains in the polymer network. A variable that received little attention is the degree of N and O acetylation of the amino sugars of the glycan chain. The analytical procedure is time-consuming, although a rapid screening method has been proposed. Membrane-

bound teichoic acid is present in all Gram-positive species, but cell wall-bound teichoic acid is present in only some Gram-positive species. Teichoic acids can easily be extracted and purified and can be analyzed by gas-liquid chromatography (101).

Cellular Fatty Acid Analysis More than 300 fatty acids and related compounds are present in bacterial cells. Polar lipids are the constituents of the lipid bilayer of bacterial membranes and have been frequently studied for classification and identification purposes. Other types of lipids, such as sphingophospholipids, occur in only a restricted number of taxa and were shown to have taxonomic value within these groups (11). Variations in chain lengths, double-bond positions, and substituent groups are very useful for the characterization of bacterial taxa (102). Mostly, the total cellular fatty acid fraction is extracted, but particular fractions, such as the polar lipids, have also been analyzed. The cellular fatty acid methyl ester composition is a stable parameter provided that highly standardized culture conditions are used. The methylated fatty acids are typically separated by gas-liquid chromatography, and both the occurrence and the relative amounts of methylated fatty acids characterize bacterial fatty acid profiles.

MALDI-TOF and Other Mass Spectrometric Methods The first reports involving the use of MALDI-TOF mass spectrometry (MS) were published in the late 1980s, and its application has increased exponentially. By now, this methodology and its many applications have revolutionized the routine practices in clinical microbiology. In MALDITOF MS, the sample is mixed with a matrix that is chosen such that it specifically absorbs a laser beam. The resulting high-energy impact is followed by the formation of ions that are extracted through an electric field and that are subsequently focused and detected as an m/z (mass/charge) spectrum. Typically, high-abundance peptides, like those derived from ribosomal protein fractions, that are of low mass and ionize readily are observed in the spectra (103). The simplicity and speed of analysis represent part of its strength, and the whole process can be highly automatized. These features make the approach particularly attractive to research laboratories that routinely deal with the analysis and identification of large numbers of bacterial isolates. In microbiology, MALDI-TOF MS is primarily being used for species-level identification of various bacteria and fungi, but its potential for infraspecific typing of bacteria is being explored (103, 104). The technique is nowadays used for an increasing range of applications, including the analysis of mixed cultures, the differentiation between antimicrobialresistant and -susceptible strains, direct identification of bacteria and yeasts in clinical specimens, and the rapid grouping of bacterial species in large collections of isolates (105). The potential of MALDI-TOF MS for bacterial identification has also been used in a number of alternative ways. These include the fast and accurate differentiation of PCR products according to their lengths and rapid analysis of PCR products and restriction fragment length polymorphism patterns of microbial samples for size determination of double-stranded amplicons and restriction fragments (106, 107). Because of limitations of these approaches by length heterogeneities of specific marker genes that diminish their discriminatory power, von Wintzingerode et al. (108)

17. Taxonomy and Classification of Bacteria n

combined base-specific cleavage of amplified 16S rRNA genes with MALDI-TOF MS. In this process, 16S rRNA gene signature sequences are amplified in the presence of dUTP instead of dTTP, followed by strand separation and uracil-DNA-glycosylase-mediated cleavage at each T-specific site. Fragment pattern detection was performed by MALDI-TOF MS and proved useful for the identification of several bacteria, including Bordetella and Mycobacterium strains (108, 109). Other studies report on the use of surface-enhanced laser desorption ionization-TOF MS for the identification of bacteria (110–112). Surface-enhanced laser desorption ionization is distinguished from MALDI in its use of an active sample probe—the ProteinChip array—that has an adsorptive surface that allows bacterial lysates to be subjected, without prior treatments, to on-chip sample preparation steps, such as selective washing and desalting. This procedure minimizes sample losses, while speeding up and simplifying sample preparation, compared to the standard methods normally employed prior to the use of MALDI. Furthermore, the active capture of the proteins by the protein chip array ensures nondiscriminatory binding of target proteins, which in turn improves the reproducibility and allows both peak mass-to-charge ratios and intensity to be used in sample characterization. Finally, the Ibis T5000 biosensor technology (Bruker Daltonics Inc., Billerica, MA) uses broad-range PCR primers that target conserved regions of bacterial genomes, such as ribosomal sequences and conserved elements from essential protein-coding genes (i.e., housekeeping genes), and is designed to rapidly detect and identify a variety of pathogens without prior knowledge of the pathogen’s nucleic acid sequence (113). The use of such broad-range priming targets across the widest possible grouping of organisms enables amplification of most species within a group. The T5000 biosensor uses electrospray ionization MS to analyze the products of broad-range PCR, which allows for the precise determination of the molecular mass of the PCR products. These high-precision mass measurements are then used to unambiguously derive base compositions of the PCR products, which are compared to a database for identifying the organism. This technology allows for a multilocus identification of bacteria in the samples with significantly less time and effort than sequencing and performs well with samples from a variety of clinical and environmental matrices, including blood, serum, various tissues, and even mosquito homogenates (114).

FTIR Spectroscopy Fourier transform infrared (FTIR) spectroscopy is used for the identification of substances in chemical analyses. In general, the wave number, the reciprocal of the wavelength, is used as the physical unit. FTIR spectroscopy involves the observation of vibrations of molecules that are excited by an infrared beam. Molecules are able to absorb the energy of distinct light quanta and start a rocking or rotation movement. An infrared spectrum represents a fingerprint that is characteristic for any chemical substance. The composition of biological material and, thus, of its FTIR spectrum is exceedingly complex, representing a characteristic fingerprint. Naumann and coworkers suggested identifying microorganisms by FTIR spectroscopy (115). In principle, a reference spectrum library is assembled based on well-characterized strains and species. The FTIR spectrum of any unidentified isolate is then measured under the same conditions as those used for the reference spectra and is compared to spectra in

263

the reference spectrum library. FTIR spectroscopy is now used for the identification and typing of a growing number of bacteria and yeasts (116–119).

Conclusions The scientifically and economically ideal identification technique remains beyond reach. Cowan’s (2) intuitive approach (which is used when the identity of the unknown is anticipated) and the stepwise method (which involves the use of dichotomous keys) suffice for numerous isolates and require only simple, rapid, and inexpensive biochemical tests. Cowan’s views are easily adapted to modern methodology. If this first-line approach fails, alternative procedures are required and available. For several reasons, including the comprehensiveness of the public databases, complete 16S rRNA gene sequence analysis is the most straightforward and obvious choice for establishing a rough identity of an isolate, although it often fails to differentiate closely related species. Much of its superiority is based on its robust capacity to reveal the phylogenetic neighborhood of the organism studied, which is information not provided by any of the other current identification protocols. This information will direct the additional analyses required for final identification to the species level. Accurate species-level identification is thus very often a two-step process in which an unknown is first assigned to a particular group, after which it can be accurately identified at the species level. The former can be achieved through sequence analysis of 16S rRNA genes using near-universal primers, the latter through an appropriate selection of housekeeping genes and specific primers once the tentative identity of an unknown is established.

NOMENCLATURE Nomenclature is the supreme generator of heat, bad temper and ill-will among taxonomists and every kind of microbiologist; the reason is that in matters of nomenclature we are all conservative. We hate change (3).

Valid Publication of Bacterial Names The International Code of Nomenclature of Bacteria (120) includes rules on how to name bacteria at different taxonomic ranks. The aim of nomenclature is to ensure that an organism is tagged with a unique name that carries valuable information. Prior to 1980, a proposal of a new bacterial taxon could be validly published in any microbiological book or journal, and the authors of the relevant sections of the successive editions of Bergey’s Manual of Determinative Bacteriology had to attempt to give a complete list of the members of any particular genus or group of genera. The unavailability of type strains and the fact that microbiologists from different disciplines were not always familiar with one another’s work caused great difficulty. All too often a worker would discover several years later that “his” or “her” organism had in fact been described earlier under a different name. To overcome such problems and others, 1 January 1980 was chosen as a new starting date for bacterial nomenclature. At that time, the Approved Lists of Bacterial Names were published on behalf of the Judicial Commission of the International Committee on Systematic Bacteriology (121). Only those names included on these lists had standing in bacterial nomenclature, and names of taxa were to be included only if they were adequately described and if a

264 n BACTERIOLOGY

type strain was available. From then onwards, all new names were validly published only in the International Journal of Systematic Bacteriology (now the International Journal of Systematic and Evolutionary Microbiology). Names can effectively be published in other journals and then validated subsequently by announcement in the Validation Lists in the International Journal of Systematic and Evolutionary Microbiology. Complete overviews of validly published names can easily be obtained through Internet sites such as http://www.bacterio.net/. One of the conditions for valid publication of names is that type strains of novel species must be deposited in two public culture collections in different countries. In case different names for the same organism are validly published, nomenclatural priority goes to the name that was validated first. As a result of this practice, all validly named species in any particular group can easily be traced and reference strains are available.

Why Do Names Change? There have been more important causes for the modification of bacterial names than the occasional detection of synonymy. As described above, our present view on bacterial classification is phylogeny based. With the advent of rRNADNA hybridization in the 1970s and subsequently of rRNA gene sequencing, taxonomists had a new framework in which they could revise classification schemes. The classical—and extreme—example is the revision of the taxonomy and nomenclature of the genus Pseudomonas, which has been proceeding painstakingly slowly over several decades. The most important reason for this slow progress is that, through the work of De Vos and De Ley (122), it became clear not only that the genus Pseudomonas consisted of five major species clusters but also that these clusters formed a polyphyletic part of a major group of bacteria now known as the Proteobacteria. Revision of the taxonomy of the pseudomonads had to consider the relationships of the various subbranches toward their numerous respective neighbors (123). The modification of our view on classification has been by far the most important reason for name changes. However, various forms of poor taxonomic practice, like the erroneous reliance on 16S rRNA similarity data rather than the level of DNA-DNA hybridization (76) or the analysis of some, rather than all, relevant species (124), also invoke a lot of changes, and hence irritation. An important reason for name changes is the lack of criteria for genus delineation. For instance, immediately after the original description of the genus Burkholderia (124), the same group of authors reclassified two of its species (B. solanacearum and B. pickettii) into the new genus Ralstonia (125). At present, the genus Burkholderia contains >70 validly named species, many of which have been isolated from soil and water samples (see chapter 43). A large phylogenetic cluster of soilborne and plant-associated Burkholderia species is considered beneficial to plants, but their classification in the genus Burkholderia hampers their exploitation as biological control or plant growth promotion agents. Although several of the opportunistic pathogens of the Burkholderia genus also have plant-beneficial properties, the phylogenetic distinctiveness of the misleadingly called “plant-beneficial group” has been used to argue for splitting the Burkholderia genus (126, 127). Also, the intrinsically inadequate descriptions of species that comprise only a single isolate, the so-called onestrain taxa (species [or genera] that are proposed on the basis of data for only one strain), have probably caused more problems than they have solved, and this is definitely

the case in the context of diagnostic microbiology. It is not possible to estimate the variability of the phenotype in the case of a species with one strain or in the case of a genus with one species and one strain, for which many recent examples exist. In diagnostic microbiology, it is well known that a species is characterized by a certain degree of variability. This variability can be measured by both phenotypic and genotypic criteria and may be revealed by simple biochemical testing or sophisticated genomic-fingerprinting techniques. In the absence of sufficient strains for quantitation of the range of divergence within a species, it will be difficult or impossible to identify new isolates of this species without DNA-DNA hybridization experiments. A classification based on results obtained with a single strain cannot be stable. Indeed, already the detection of a second strain will inevitably necessitate revision of the original species description. As a concluding remark, it should be mentioned that there is no “undo” function in bacterial nomenclature. A name that was validly published remains valid regardless of the number of modifications it undergoes thereafter. For instance, the changes of the name Pseudomonas maltophilia to Xanthomonas maltophilia and finally to Stenotrophomonas maltophilia (128) may be reasonable to some taxonomists, but the changes, particularly the most recent, have been refuted by many clinical microbiologists. As these three names were all proposed according to the rules of bacterial nomenclature, they were all validated, and the use of each of them is correct and valid.

CONCLUSIONS There seems to be a clear and bright future for bacterial taxonomy in the genomics era. A much broader range of taxonomic studies of bacteria has gradually replaced the former reliance on morphological, physiological, and biochemical characterization. This polyphasic taxonomy takes into account all available phenotypic and genotypic information and integrates it in a consensus type of classification, framed in a general phylogeny derived from 16S rRNA gene sequence analysis. The bacterial species appears as a group of isolates that originated from a common ancestor population in which a steady generation of genetic diversity resulted in clones that had different degrees of recombination and that were characterized by a certain degree of phenotypic consistency, a significant degree of DNA-DNA hybridization, and a high degree of 16S rRNA gene sequence similarity (9). Whole-genome sequences will make the practice of polyphasic taxonomy even more complete, and several genomic parameters like pcDNA, MUM index, and ANI values are ready to replace DNA-DNA hybridization as the cornerstone of species description. While the old DNA-DNA hybridization species threshold is being replaced by MLSA or ANI value-based thresholds (5, 13, 23, 44), a fair reappraisal of bacterial taxonomy and the species definition will require more than a mere methodological translation of threshold levels. Several considerations should be made. First, whole-genome sequence studies confirm that in the space of microbial diversity many cores can be distinguished (see, e.g., reference 43), and these cores very often correspond with established species that are phenotypically but not necessarily ecologically coherent. It would be wrong to let the discussion be steered by an attempt to hold on to species boundaries defined by DNADNA hybridization experiments. A new discussion on the bacterial species concept, directed by insights in genome

17. Taxonomy and Classification of Bacteria n

evolution and metagenomics but also by practical concerns (classification should remain practical and facilitate identification), is badly needed (18, 33, 129) and should yield whole-genome sequence-based thresholds, irrespective of their degree of correlation with DNA-DNA hybridization thresholds. A second key point in the discussion of a modern bacterial species definition was raised by Sutcliff et al. (10), who called for a significant reappraisal of the procedures used to describe novel prokaryotic taxa, including the introduction of new publication formats. They rightly pointed out that although progress in the description of new microbial taxa is being made at accelerating rates, there is an enormous backlog of work and conservative estimates suggest that the presently described bacterial species (0.5-mm colonies) streptococci of the pyogenic group. Species from the pyogenic or beta-

doi:10.1128/9781555817381.ch22

383

384 n

BACTERIOLOGY

TABLE 1

Phenotypic characteristics of beta-hemolytic streptococcia

Species

Lancefield group(s)

Colony sizee

Hostsi

Bacitracin susceptibility

PYRf

Humans Humans, cows Animals

+ −

+ −

− +

− −

− +

+ v

− −

−

−

−

−

−

+

v

CAMPg VPh

Hippurate Trehalose hydrolysis

Sorbitol

S. pyogenes S. agalactiae

A B

Large Large

S. dysgalactiae subsp. dysgalactiaeb S. dysgalactiae subsp. equisimilis S. equi subsp. equi S. equi subsp. zooepidemicusc S. canisc

C

Large

A, C, G, L

Large

Humans (animals)

−

−

−

−

−

+

−

C

Large

Animals

−

−

−

−

−

−

−

C

Large

−

−

−

−

−

−

+

G

Large

−

−

+

−

−

v

−

S. anginosus groupd S. porcinusc

A, C, G, F, none E, P, U, V, none

Small

Animals (humans) Dogs (humans) Humans

−

−

−

+

−

+

−

Swine (humans)

−

+

+

+

v

+

+

Large

a

Symbols and abbreviations: +, positive; −, negative; v, variable. S. dysgalactiae subsp. dysgalactiae is alpha-hemolytic on sheep blood agar plates. S. equi subsp. zooepidemicus, S. canis, and S. porcinus are primarily animal pathogens that are only rarely isolated from humans. d Species included in the S. anginosus group can be beta-hemolytic, alpha-hemolytic, or nonhemolytic on sheep blood agar plates. e Size: large, >0.5 mm after 24-h incubation; small, 0.5-mm colony) beta-hemolytic streptococci of the pyogenic group. Association of certain species with specific isolation sites has been reported. While S. anginosus is frequently found in specimens from the urogenital or gastrointestinal tracts, S. constellatus is commonly isolated from the respiratory

22. Streptococcus n 387

tract, and S. intermedius is most often identified in abscesses of the brain or liver.

Streptococcus salivarius Group Streptococcal species that belong to the S. salivarius group include S. salivarius and S. vestibularis. They have been primarily isolated from the oral cavity and blood. Another species of this group, S. thermophilus, is found only in dairy products. S. salivarius has been repeatedly reported as a cause of bacteremia, endocarditis, and meningitis (sometimes iatrogenic), while S. vestibularis has not been clearly associated with human infection. Isolation of S. salivarius from blood cultures does correlate to some extent with neoplasia (49, 50).

Streptococcus mutans Group S. mutans and S. sobrinus belong to the S. mutans group. They are the most commonly isolated species of the group that originate from human clinical specimens, usually obtained from the oral cavity. S. criceti, S. ratti, and S. downei have occasionally been identified from human sources, while the other streptococcal species of the S. mutans group (S. ferus, S. macacae, S. hyovaginalis, and S. devriesei) have been identified only in animals. S. mutans is the primary etiologic agent of dental caries, and infection is transmissible. By 18 years of age, 85% of the population have at least one carious lesion (51). Permanent colonization with S. mutans occurs under normal living conditions in the Western world between the second and the end of the third year of life (51). Molecular analysis of mother and infant isolates reveals that strains are usually acquired from the mother and that the colonization rate of infants depends on the bacterial load of the mother (52). Analyses of streptococcal blood culture isolates show that S. mutans is the most frequently isolated species of this group in cases of bacteremia (1).

Streptococcus bovis Group Extensive taxonomic changes have occurred in this group, and strains formerly known as human S. bovis isolates are designated as different species (see “Taxonomy” above). The group now includes S. equinus, S. gallolyticus, S. infantarius, and S. alactolyticus. Species from this group are frequently encountered in blood cultures of patients with bacteremia, sepsis, and endocarditis. The clinical significance of blood cultures growing streptococci from the S. bovis group lies in the association of (i) S. gallolyticus subsp. gallolyticus with gastrointestinal disorders, including colon cancer and chronic liver disease, and (ii) S. gallolyticus subsp. pasteurianus with meningitis (53–55).

Other Streptococci Infrequently Isolated from Human Specimens Streptococcal species that are primarily animal pathogens are sometimes isolated from human hosts, in most cases from humans that are in close contact with animals. S. suis, S. porcinus, and S. iniae belong to this category. S. suis is a swine pathogen that has occasionally been isolated from cases of human meningitis and bacteremia. S. suis is encapsulated and appears to be alpha-hemolytic on sheep blood agar plates, although some strains are beta-hemolytic on horse blood agar. S. suis strains are positive for the Lancefield group antigen R, S, or T, which helps to distinguish them from the phenotypically similar species S. gordonii, S. sanguinis, and S. parasanguinis. Similar to S. suis, S. porcinus (Lancefield groups E, P, U, and V) is primarily a swine pathogen. Beta-hemolytic S. porcinus strains have rarely

been isolated from human sources such as peripheral blood, wounds, and the female genital tract (56). Molecular studies of S. porcinus isolates from the female genital tract, however, indicate that these isolates belong to a novel species designated S. pseudoporcinus (57). S. pseudoporcinus may be more prevalent in the female genital tract than previously assumed (58). Strains can easily be misidentified as S. agalactiae due to isolation from the female genital tract, false-positive reactions with commercially available group B antisera, and a positive CAMP test reaction. S. porcinus and S. pseudoporcinus can be L-pyrrolidonyl-beta-naphthylamide (PYR) positive and do not hydrolyze hippurate, in contrast to S. agalactiae. S. iniae is a fish pathogen that is beta-hemolytic but does not possess any Lancefield group antigens. It has been isolated from soft tissue infections, bacteremia, endocarditis, and meningitis in people handling fish (59, 60). S. iniae isolates resemble S. pyogenes strains due to the fact that both are PYR positive. Beta-hemolysis of the species can be observed only around agar stabs or under anaerobic culture conditions. Commercial identification systems do not correctly identify the species; the failure to react with Lancefield group antisera is important to notice, since it is rare among beta-hemolytic streptococci. Very recently, a novel streptococcal species associated with the handling of fish has been reported as a human pathogen. The species has been designated S. hongkongensis; it is closely related to S. iniae and S. parauberis (61).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS Specimens suspected of harboring streptococci should be collected by the methods outlined elsewhere in this Manual (chapter 18). Since many streptococcal species lose viability fairly quickly, it is best to place swabs in an appropriate moist transport medium and process specimens rapidly. If transport time is below 1 to 2 h, a special transport system is not absolutely necessary. S. pyogenes can safely be transported on dry swabs; desiccation enhances recovery from mixed cultures by suppression of the accompanying microbiota (62). Detailed recommendations for collection and storage of swabs from pregnant women to detect S. agalactiae colonization have been issued by the U.S. Centers for Disease Control and Prevention (35). These recommendations are summarized below under “Special Procedures for Streptococcus agalactiae Screening.”

DIRECT EXAMINATION Microscopy Microscopic examination shows streptococci as Grampositive bacteria growing in chains of various lengths. S. pneumoniae isolates most often present as Gram-positive diplococci with an elongated appearance, but a reliable microscopic distinction of S. pneumoniae from enterococci and other streptococci is not possible. In blood culture specimens, S. pneumoniae tends to form chains of varying lengths, similar to other streptococci. Direct identification of streptococci by microscopic methods is most helpful in the case of clinical specimens from sterile body sites, such as cerebrospinal fluid. Tiny, irregular cocci in clumps of chains seen in abscess- or peritonitis-associated aspirates are suggestive of the S. anginosus group. Interpretation of Gram stain results from nonsterile body sites is difficult due to the residential microbiota, which frequently includes streptococci. Thus, for example, throat

388 n

BACTERIOLOGY

swabs should not be examined by Gram stain for diagnosis of “strep” throat.

Direct Antigen Detection of S. pyogenes from Throat Specimens S. pyogenes is the most common cause of acute pharyngitis and accounts for 15 to 30% of cases of acute pharyngitis in children and 5 to 10% of cases in adults. If the diagnosis can be provided rapidly, antibiotic therapy can be initiated promptly to relieve symptoms, to avoid sequelae, and to reduce transmission. Numerous assays for direct detection of the group A-specific carbohydrate antigen in throat swabs by agglutination methods or immunoassays (enzyme, liposome, or optical), also referred to as “rapid antigen assays,” have become commercially available during the past 2 decades. A list of FDA-cleared tests is accessible via the Internet (http://www.accessdata.fda.gov/scripts/cdrh/devices atfda/index.cfm?Search_Term=866.3740). Although these tests provide rapid results and allow early treatment decisions, the throat culture remains the gold standard. Sensitivities of rapid antigen tests range from 70% to 96% and have never equaled that of culture (63, 64). Negative rapid antigen test results should therefore be confirmed by culture in children and adolescents when typical clinical signs are present (65). In adults, the confirmation of negative rapid antigen test results has not been regarded as necessary for many years, but the most recent Infectious Diseases Society of America (IDSA) guidelines challenge this concept (66). The specificity, however, is generally high, even though false-positive antigen results are seen from patients previously diagnosed and/or treated for S. pyogenes (67). Moreover, the low positive predictive value of rapid group A antigen tests in the adult population frequently results in prescribing unnecessary antimicrobial therapy (68).

Antigen Detection of S. agalactiae in Urogenital Tract Samples Several different commercially available antigen detection tests have been developed for the identification of S. agalactiae in samples from the urogenital tract. Independent from the technique involved (latex agglutination, enzyme immunoassay, or optical immunoassay), without a prior cultural enrichment step, all of the currently available tests lack sufficient sensitivities to detect bacterial colonization with S. agalactiae (69). They are not recommended for screening of pregnant women by the CDC (35).

Antigen Detection of S. pneumoniae in Urine Samples An immunochromatographic membrane test relying on the detection of the cell wall-associated polysaccharide that is common to all S. pneumoniae serotypes (C-polysaccharide antigen) (Binax NOW; Binax Inc., Portland, ME) has proven helpful for the identification of S. pneumoniae infections in adult patients, especially in patients that already received antibiotic treatment. In contrast to conventional diagnostic methods, reported sensitivities of antigen detection in urine samples range between 50 and 80%, and specificities are higher than 90% (70, 71). Following pneumococcal infection, the test can remain positive for about 1 to 6 months (72). Due to the fact that the test is also positive in S. pneumoniae carriage without infection, as is often observed among infants (73), it is of limited value in pediatric patients. The test should not be used in children below the age of 6 (73), and comprehensive studies on schoolchildren with lower colonization rates have not been performed.

It can currently be recommended only in adults as an addition to conventional diagnostic culture techniques for S. pneumoniae (74) and is probably most helpful in patients who received antimicrobial treatment before cultures were obtained.

Streptococcal Antigen Detection in CSF Commercially available antigen detection tests for the diagnosis of pathogenic microorganisms in cerebrospinal fluid (CSF) samples include reagents for the detection of S. agalactiae and S. pneumoniae. These tests have also been used on positive blood culture specimens. The tests are not recommended for routine use, as the results should not be used to change decisions about empiric therapy based on clinical and laboratory criteria (75). It has also been shown that the sensitivity of direct antigen detection in CSF is low (0.5 mm after 24 h of incubation, in

contrast to beta-hemolytic strains of the S. anginosus group (formerly called the “S. milleri” group), which present with pinpoint colonies of ≤0.5 mm after the same incubation time (Fig. 1). Members of the S. anginosus group emit a distinct odor resembling butterscotch or caramel, presumably due to the production of diacetyl by the species belonging to this group. Among the beta-hemolytic species of the pyogenic group, S. agalactiae produces the largest colonies with a relatively small zone of hemolysis. Nonhemolytic S. agalactiae strains do occur and resemble enterococci. Within the group of alpha-hemolytic streptococci, S. pneumoniae has a colony morphology that helps to distinguish pneumococcal isolates from other streptococci of the viridans group. Due to the production of capsular polysaccharide, colonies glisten and appear moist. Colonies may be large and mucoid if large amounts of capsular polysaccharide are made, a feature often encountered in serotype 3

FIGURE 1 Colony morphology of selected streptococci. (A) S. pyogenes strain ATCC12344; (B) clinical isolate of S. agalactiae; (C) clinical isolate of S. dysgalactiae subsp. equisimilis (Lancefield group G); (D) mixed culture of S. anginosus (ATCC12395, open arrow) and clinical isolate of S. dysgalactiae subsp. equisimilis (closed arrow) (note the difference in colony size); (E) clinical isolate of S. pneumoniae (note the central depression of the colonies); (F) clinical isolate of S. pneumoniae (note the mucoid appearance of colonies). doi:10.1128/9781555817381.ch22.f1

22. Streptococcus n 391

strains. This phenotype is usually typical for S. pneumoniae but can also occasionally be observed in S. pyogenes. Another characteristic feature of S. pneumoniae is the central navel-like depression of the colonies that is caused by the pneumococcal autolysin. Other viridans group streptococci lack this feature and have a dome-like appearance; however, up to 20% of S. pneumoniae strains display a phenotype that is indistinguishable from that of viridans group streptococci (96). Nonhemolytic gray colonies are typical for species of the S. bovis and S. salivarius groups. Typical streptococcal colony morphologies are presented in Fig. 1.

PYR testing, and strains of other related genera may be PYR positive (including the genera Abiotrophia, Aerococcus, Enterococcus, Gemella, and Lactococcus). However, PYR-positive beta-hemolytic enterococcal isolates typically present with a different colonial morphology (smaller zone of betahemolysis and bigger colony size) and, when combined with other phenotypic characteristics (see chapter 23), may be distinguished from streptococci. To avoid false-positive reactions caused by other PYR-positive bacterial species (for example, staphylococci), the test should be performed only on pure cultures.

Identification of Beta-Hemolytic Streptococci by Lancefield Antigen Immunoassays

Bacitracin Susceptibility

Commercially available Lancefield antigen grouping sera are primarily used for the differentiation of beta-hemolytic streptococci. Products for rapid antigen extraction and subsequent agglutination can be obtained from many different suppliers. The presence of the Lancefield group B antigen in beta-hemolytic isolates from human clinical specimens correlates with the species S. agalactiae, but cross-reactivity of the group B antigen with the newly described species S. pseudoporcinus has been reported (58). Similarly, the detection of the Lancefield group F antigen in small-colonyforming streptococci from human clinical material allows a fairly reliable identification of a strain as a member of the S. anginosus group. The presence of Lancefield group A, C, or G antigens necessitates further testing (Table 1). Betahemolytic streptococcal strains not reacting with any of the Lancefield antisera are rare and should be further identified by phenotypic tests or nucleic acid detection techniques.

With rare exceptions, S. pyogenes displays bacitracin susceptibility, in contrast to other human beta-hemolytic streptococci. Together with Lancefield antigen determination, it can be used for the identification of S. pyogenes, since betahemolytic strains of other streptococcal species that may contain the group A antigen are bacitracin resistant. The test can also be used to distinguish S. pyogenes from other PYR-positive beta-hemolytic streptococci (S. iniae and S. porcinus). A bacitracin disk (0.04 U) is applied to a sheep blood agar (SBA) plate that has been heavily inoculated with 3 or 4 colonies of a pure culture of the strain to be tested. It is important to perform the test from a subculture on SBA, since placement of bacitracin disks on primary plates is not sensitive enough. After overnight incubation at 35°C in 5% CO2, any zone of inhibition around the disk is interpreted as indicating susceptibility. Importantly, bacitracin-resistant S. pyogenes isolates have been reported and clusters of bacitracin-resistant strains were observed in several European countries (98–100).

Identification of Beta-Hemolytic Streptococci with Phenotypic Tests and MALDI-TOF MS

VP Test

A number of streptococcal identification products incorporating batteries of physiologic tests are commercially available (see chapters 4 and 19). In general, these products perform well with commonly isolated pathogenic streptococci but may lack accuracy for identifying streptococci of the viridans group. The MALDI-TOF (Bruker Daltonics, Billerica, MA)-based bacterial identification also has limitations concerning several streptococcal species. While among beta-hemolytic streptococci, identification of S. pyogenes and S. agalactiae corresponds well to conventional tests, the correct identification of S. dysgalactiae to the species level cannot be achieved for the majority of isolates (97). Further problems include the misidentification of S. mitis or S. oralis as S. pneumoniae and continuing difficulties in the correct species identification of viridans group streptococci. For the bulk of pathogenic streptococci isolated in clinical laboratories (e.g., S. pyogenes, S. agalactiae, and S. pneumoniae), serologic or presumptive physiologic tests (as described below) still offer an acceptable alternative to commercially available identification systems.

PYR Test The presence of the enzyme pyrrolidonyl aminopeptidase is often tested to distinguish S. pyogenes from other betahemolytic streptococci. Hydrolysis of L-pyrrolidonyl-betanaphthylamide by the enzyme to β-naphthylamide produces a red color with the addition of cinnamaldehyde reagent (see chapter 19). The beta-hemolytic streptococcal species S. iniae, S. porcinus, and S. pseudoporcinus can be PYR positive but are only rarely identified in human clinical specimens. PYR spot tests are commercially available. It is important to distinguish Streptococcus from Enterococcus prior to

The Voges-Proskauer (VP) test detects the formation of acetoin from glucose fermentation. It is performed on streptococci as a modification of the classical VP reaction that is used for the differentiation of enteric bacteria. Smallcolony-forming beta-hemolytic streptococci of the S. anginosus group that are VP positive may be distinguished from large-colony-forming beta-hemolytic streptococci harboring identical Lancefield antigens (A, C, or G). Streptococci of the S. mitis group are VP negative. For the modified VP reaction as described by Facklam and Washington in the 5th edition of this Manual (160), the culture growth of an entire agar plate is used to inoculate 2 ml of VP broth and incubated at 35°C for 6 h. Following the addition of 5% α-naphthol and 40% KOH, the tube is shaken vigorously for a few seconds and incubated at room temperature for 30 min. A positive test yields a pink-red color that results from the reaction of diacetyl with guanidine.

BGUR Test

Detection of β-glucuronidase (BGUR) activity distinguishes S. dysgalactiae subsp. equisimilis strains containing Lancefield group antigen C or G from BGUR-negative, small-colonyforming streptococci of the S. anginosus group with the same Lancefield group antigens. Rapid methods for the BGUR test are commercially available. Alternatively, a rapid fluorogenic assay with methylumbelliferyl-β- D -glucuronide (MUG)-containing MacConkey agar, often used for Escherichia coli, has been described (101).

CAMP Test The CAMP factor reaction was first described in 1944 by Christie, Atkins, and Munch-Petersen and refers to the synergistic lysis of erythrocytes by the beta-hemolysin of

392 n

BACTERIOLOGY

serving the development of a deep purple color, signifying a positive test (103). Streptococci other than S. agalactiae may also be hippurate hydrolysis positive, especially viridans group streptococci.

Identification of S. pneumoniae and Viridans Group Streptococci

FIGURE 2 CAMP factor test. Arrowhead-shaped zone of hemolysis in the zone of the S. aureus beta-hemolysin. (A) Clinical isolate of a weakly beta-hemolytic S. agalactiae strain. (B) Beta-hemolytic S. agalactiae strain O90R. (C) Nonhemolytic S. agalactiae strain R268. doi:10.1128/9781555817381.ch22.f2

Staphylococcus aureus and the extracellular CAMP factor of S. agalactiae. The gene (cfb) and its expression can be demonstrated in the vast majority (>98%) of S. agalactiae isolates, but CAMP-negative mutants do occur. The strain to be tested and a Staphylococcus aureus strain (ATCC 25923) are streaked onto a sheep blood agar plate at a 90° angle. Plates are incubated in ambient air overnight at 36 ± 1°C. A positive reaction can be detected by the presence of a triangular zone of enhanced beta-hemolysis in the diffusion zone of the beta-hemolysin of S. aureus and the CAMP factor (Fig. 2). CAMP factor-positive strains can also be detected by a method using β-lysin-containing disks (Remel Inc., Lenexa, KS) or by a rapid CAMP factor spot method (102). Despite the fact that close homologs of the CAMP factor gene are present in many S. pyogenes strains, most beta-hemolytic streptococci other than S. agalactiae are negative in the above-described CAMP factor test, except for the rare human isolates of S. iniae, S. porcinus, and S. pseudoporcinus. Several Gram-positive rods including corynebacteria and Listeria monocytogenes strains may also be CAMP factor positive.

Hippurate Hydrolysis Test The ability to hydrolyze hippurate is an alternative test for the presumptive identification of S. agalactiae. A rapid version of the test, as it is used for campylobacters, can be performed by incubating a turbid suspension of bacterial cells in 0.5 ml of 1% aqueous sodium hippurate for 2 h at 35°C. Glycine formed as an end product of hippurate hydrolysis is detected by adding ninhydrin reagent and ob-

The correct species identification of viridans group streptococci other than S. pneumoniae is challenging. Recent taxonomic changes and identification of novel streptococcal species have further complicated matters. The number of recognized species in this group is now greater than 30. The viridans group includes alpha-hemolytic, nonhemolytic (S. salivarius group and S. bovis group), and beta-hemolytic (S. anginosus group) streptococcal strains. All of the viridans group streptococci are leucine aminopeptidase positive and pyrrolidonyl arylamidase negative. Conventional microbiologic tests are limited with respect to species identification but are helpful in placing isolates into the correct streptococcal groups (Table 2). Beighton et al. described an identification scheme based on phenotypic tests that allowed the differentiation and correct species identification of the majority of viridans group species (104). The scheme requires the evaluation of enzymatic reactions performed by in-house fluorogenic tests that are not commercially available. Importantly, most clinical laboratories must strive for group, instead of species, classifications with current phenotypic test panels. The API tests (bioMérieux, Marcy l’Etoile, France) offer species identification of viridans group streptococci. While many species from this group are identified with acceptable accuracy, several species have not been included in the database. Comparisons of molecular species identification by DNA reassociation studies with the results of the API Rapid ID 32 Strep system showed that more than 85% of 156 strains from streptococcal species included in the database were correctly identified (105). However, for species not included in the database, more than 50% were incorrectly identified by the test (105). Evaluation studies performed under routine clinical conditions appear to yield less favorable results (106). Evaluations of the Vitek 2 (bioMérieux) automated phenotypic identification system showed that streptococcal group assignment for 75% of isolates were concordant with 16S rRNA gene sequencing data (107). In another study, correct identification at the species level was accomplished for only 55% of the tested isolates (108). Similarly, some publications on MALDI-TOF MS technology report major problems with the correct species identification of streptococci from the viridans group and a frequent misidentification of S. pneumoniae (97, 109). However, very recent publications indicate that a different system (Vitek MS; bioMérieux) has solved this problem for S. pneumoniae (110, 111). Novel commercial identification systems for streptococci include the FDA-cleared Verigene Grampositive blood culture (BC-GP) nucleic acid test (Nanosphere, Inc., Northbrook, IL) and the FilmArray platform (FA; BioFire, Salt Lake City, UT) for the identification of bacterial pathogens directly from blood culture bottles (112, 113). While very promising results are published for major bacterial pathogens, including S. pyogenes and S. agalactiae, a reliable species identification for S. pneumoniae and streptococci from the viridans group is currently not possible with these tests. In conclusion, phenotypic and automated species identification of viridans group streptococci remains challenging, and acceptable results can currently be achieved only at the group level.

22. Streptococcus TABLE 2

mitisb anginosusc mutansd salivariuse bovisf

393

Phenotypic characteristics of major streptococcal groupsa

Streptococcal group S. S. S. S. S.

n

Arginine hydrolysis

Esculin

Mannitol

Sorbitol

Urea hydrolysis

VP

v + − − −

v + + v v

− − + − v

v − + − −

− − − v −

− + + + +

a

Symbols and abbreviations: +, positive; −, negative; v, variable. The S. mitis group comprises the species S. mitis, S. sanguinis, S. parasanguinis, S. gordonii, S. cristatus, S. oralis, S. infantis, S. peroris, S. australis, S. sinensis, S. orisratti, S. oligofermentans, and S. massiliensis. S. sanguinis, S. parasanguinis, S. gordonii, and S. cristatus are arginine hydrolysis positive; other species from the S. mitis group are arginine hydrolysis negative. c S. anginosus, S. constellatus, and S. intermedius belong to the S. anginosus group. d The S. mutans group includes S. mutans, S. sobrinus, and the following species rarely isolated from humans: S. criceti, S. ratti, and S. downei. e The S. salivarius group contains S. salivarius, S. vestibularis, and S. thermophilus. S. salivarius is variable, S. vestibularis is positive, and S. thermophilus is negative for urea hydrolysis. f The S. bovis group now includes S. equinus, S. gallolyticus, S. infantarius, and S. alactolyticus. S. gallolyticus subsp. gallolyticus is positive for the acidification of mannitol, and the other species from the S. bovis group are negative. b

Molecular methods may offer alternative approaches to conventional phenotypic identification schemes. The most common molecular identification method, 16S rRNA gene sequencing, does not yield reliable species identification for several species including S. mitis, S. oralis, and S. pneumoniae. The 16S rRNA gene sequences are more than 99% identical (4). Sequence determination of the manganese-dependent superoxide dismutase gene sodA appears to be more reliable (108, 114). In contrast to 16S rRNA sequencing, it allows the differentiation of S. mitis, S. oralis, and S. pneumoniae and the correct identification of streptococcal species from the viridans group. Descriptions of the species belonging to the viridans group streptococci are given below, and physiological traits of the groups are shown in Table 2.

S. mitis Group The large number of different streptococcal species belonging to the S. mitis group has been mentioned earlier. This group of predominantly alpha-hemolytic streptococci includes several species of known clinical significance together with others for which few or no clinical data have been collected. Among the phenotypic characteristics of the species in this group, extracellular polysaccharide production is negative for S. mitis strains but is a variable characteristic of S. oralis isolates. This feature correlates with the smooth colony surface of many S. oralis strains and the rough and dry appearance of S. mitis colonies.

S. anginosus Group The small-colony-forming species S. anginosus, S. constellatus, and S. intermedius belong to the S. anginosus group. Strains of the S. anginosus group may be non-, alpha-, or beta-hemolytic on blood agar plates with some variations between the species. While S. constellatus is frequently betahemolytic, most isolates of S. intermedius are nonhemolytic. For many strains, growth is enhanced in the presence of CO2 with some strains requiring anaerobic conditions. S. anginosus and S. constellatus strains may possess Lancefield group antigen A, C, F, or G. Most S. constellatus or S. intermedius strains react with antisera against Lancefield group F antigen or are nongroupable. The species S. constellatus has been further subdivided into two subspecies, S. constellatus subsp. constellatus and S. constellatus subsp. pharyngis (115). S. constellatus subsp. constellatus is phenotypically different from S. constellatus

subsp. pharyngis, which usually possesses the Lancefield group antigen C, is beta-hemolytic, and has been associated with pharyngitis. Detailed phenotypic characteristics of the S. anginosus group are shown in Table 3 (1, 116, 117).

S. mutans Group The S. mutans group includes S. mutans, S. sobrinus, S. criceti, S. ratti, S. downei, S. ferus, S. hyovaginalis, S. devriesei, and S. macacae. S. mutans and S. sobrinus are frequently found in human hosts, while the other species are only rarely encountered in humans or represent animal pathogens. The species of the S. mutans group are characterized by the production of extracellular polysaccharides from sucrose, which can be tested by culturing the bacteria on sucrosecontaining agar and by the ability to produce acid from a relatively wide range of carbohydrates. S. mutans strains may present with an atypical morphology for streptococci, forming short rods on solid medium or in broth culture under acidic conditions. On blood agar, colonies are often hard and adherent and usually alpha-hemolytic. Under anaerobic growth conditions, some strains are beta-hemolytic. S. sobrinus strains are mostly nonhemolytic or occasionally alpha-hemolytic. On sucrose-containing agar, species from this group form colonies that are rough (frosted-glass appearance), heaped, and surrounded by liquid-containing glucan.

S. salivarius Group Streptococcal species in the S. salivarius group are S. salivarius, S. vestibularis, and S. thermophilus. S. salivarius strains are usually non- or alpha-hemolytic on blood agar. On sucrose-containing agar, strains form large mucoid or hard colonies due to the production of extracellular polysaccharides. A high proportion of S. salivarius strains react with the Lancefield group K antiserum. Species in this group may also react with the streptococcal group D antiserum. It is unclear if these strains truly possess the group D antigen or yield a nonspecific cross-reaction. S. vestibularis is alphahemolytic, and the failure of this species to produce extracellular polysaccharides on sucrose-containing agar is helpful in distinguishing S. vestibularis from S. salivarius strains. S. thermophilus is found in dairy products but has not been isolated from clinical specimens.

S. bovis Group The species belonging to the S. bovis group (S. equinus, S. gallolyticus, S. infantarius, and S. alactolyticus) are nonentero-

394 n

BACTERIOLOGY TABLE 3

Phenotypic characteristics of streptococcal species from the S. anginosus groupa S. anginosus

S. constellatusb

S. intermedius

β-D-Fucosidase

−

−

+

β-N-Acetylglucosaminidase

−

−

+

β-N-Acetylgalatosaminidase

−

−

+

Neuraminidase

−

−

+

α-D-Glucosidase

v

+

+

β-D-Glucosidase

+

−

v

β-D-Galactosidase

v

−

+

Amygdalin (acidification)

+

v

v

Mannitol (acidification)

v

−

−

Sorbitol (acidification)

−

−

−

Lactose (acidification)

+

v

+

Arginine hydrolysis

+

+

+

Esculin hydrolysis

+

v

+

VPc

+

+

+

Urease

−

−

−

Test

a Symbols and abbreviations: +, positive; −, negative; v, variable; data from Summanen et al. (116), Facklam (1), and Whiley (117). b The species S. constellatus subsp. pharyngis is β-D-fucosidase, β-D-acetylglucosaminidase, β-D-acetylgalactosaminidase, and β-D-glucosidase positive, in contrast to S. constellatus subsp. constellatus. c Voges-Proskauer test (formation of acetoin from glucose fermentation).

coccal group D streptococci that are PYR negative. Most strains grow on bile esculin agar and are unable to grow in 6.5% NaCl. On blood agar, strains are either nonhemolytic or alpha-hemolytic. Strains of the S. bovis group share phenotypic characteristics with S. mutans strains, such as production of glucan, fermentation of mannitol, and growth on bile esculin agar. However, the S. bovis group does not ferment sorbitol. S. gallolyticus subsp. gallolyticus and S. infantarius subsp. coli typically ferment starch or glycogen and give a Lancefield group D reaction, in contrast to S. gallolyticus subsp. pasteurianus. A detailed phenotypic characterization and emended description of the different subspecies has recently been published (53). For the identification of species in this group, testing of β-glucuronidase, α- and βgalactosidase, β-mannosidase, acid production from starch, glycogen, inulin, and mannitol is helpful. As described, strains formerly known as S. bovis currently belong to several species of the S. bovis group.

Physiologic Tests Optochin Test Most S. pneumoniae isolates are optochin susceptible. Before application of the optochin disk, several colonies of a pure culture are streaked on a sheep blood agar plate. Optochin testing should be performed on plates that are incubated at 35 to 36°C overnight in 5% CO2 because up to 8% of strains do not grow under ambient conditions. S.

pneumoniae isolates show zones of inhibition of ≥14 mm with a 6-mm disk (containing 5 μg of optochin) and zones of inhibition of ≥16 mm with a 10-mm disk. Incubation in 5% CO2 yielded increased specificity (6, 96). Optochinresistant S. pneumoniae strains have been reported as well as optochin-susceptible S. mitis isolates (especially when tested under ambient conditions). Since optochin testing may miss between 4% and 11% of bile-soluble S. pneumoniae isolates (6, 96), strains displaying a smaller zone of inhibition (9 to 13 mm for the 6-mm disk) should be subjected to additional testing (e.g., bile solubility or genetic testing) to confirm species identification. Application of an optochin disk onto the primary culture medium may facilitate a rapid presumptive identification but may miss pneumococcal isolates in a mixed culture. The optochin susceptibility test should be repeated with a pure culture in cases of mixed cultures, or additional tests should be performed (e.g., bile solubility).

Bile Solubility Test The bile solubility test can be performed either in a test tube or by direct application of the reagent to an agar plate. For the test tube method, a saline suspension of a pure culture is adjusted to a McFarland standard of 0.5 to 1.0, and 0.5 ml of the suspension is added to a small tube. The bacterial suspension is mixed with 0.5 ml of 10% sodium deoxycholate (bile) and incubated at 35°C. A control containing 0.5 ml of bacterial suspension with 0.5 ml of saline

22. Streptococcus

should be prepared for each strain tested. A positive result is characterized by clearing of the bile suspension within 3 h; clearing can start as early as 5 to 15 min after inoculation and allows the identification of a strain as S. pneumoniae. For the plate method, 1 drop of 10% sodium deoxycholate is placed directly on a colony of the strain in question, and the plate is incubated at 35°C for 15 to 30 min in ambient air. It is important to keep the plates in a horizontal position in order to prevent the reagent from washing away the colony. Colonies of S. pneumoniae disappear or demonstrate a flattened colony morphology, while other viridans group streptococci appear unchanged. In contrast to optochin susceptibility testing, bile solubility testing demonstrated excellent sensitivity and specificity in a recent comprehensive evaluation (96).

n

395

Esculin Hydrolysis Esculin agar slants (Becton Dickinson and other sources) are inoculated and incubated for up to 1 week. A positive reaction appears as a blackening of the medium; no change in color indicates a negative esculin hydrolysis test result.

Hyaluronidase Production Hyaluronidase activity can be detected on brain heart infusion agar plates supplemented with 2 mg/liter of sodium hyaluronate (Sigma-Aldrich, St. Louis, MO). The strains to be tested are inoculated by stabbing into the agar, and plates are incubated anaerobically at 37°C overnight. After the plate is flooded with 2 M acetic acid, hyaluronidase activity is indicated by the appearance of a clear zone around the stab. A quantitative method for determining hyaluronidase activity can be performed in microtiter trays (120).

Bile Esculin Test Production of Extracellular Polysaccharide For the bile esculin test, bile esculin medium (available from commercial sources) in either plates or slants should be inoculated with one to three colonies of the organism to be tested and incubated at 35°C in ambient air for up to 48 h. Optimal results can be achieved by using medium supplemented with 4% oxgall (the equivalent of 40% bile) (Remel, Lenexa, KS) and a standardized inoculum of 106 CFU (118). A definitive blackening of plated medium or blackening of at least one-half of an agar slant is considered a positive result, indicative of species belonging to the S. bovis group or enterococci. Occasional other viridans group streptococci are positive with this test or display weakly positive reactions that are difficult to interpret. Isolates from patients with serious infections (e.g., endocarditis) should be more completely characterized.

Arginine Hydrolysis Arginine hydrolysis is a key reaction for the identification of viridans group streptococci. Discrepancies can occur among test methods (119). Two commonly used methods are detailed here. Moeller’s decarboxylase broth containing arginine (Becton Dickinson, Franklin Lakes, NJ, and other sources) should be inoculated with the test organism, overlaid with mineral oil, and incubated at 35 to 37°C for up to 7 days. Degradation of arginine results in elevated pH, indicated by development of a purple color. Negative results are indicated by a yellow color, which is due to acid accumulation from metabolism of glucose only. For the microtiter plate method (104), 3 drops of the arginine-containing reagent are inoculated with 1 drop of an overnight ToddHewitt broth culture and incubated for 24 h at 37°C anaerobically. Production of ammonia is detected by the appearance of an orange color following addition of 1 drop of Nessler’s reagent.

Urea Hydrolysis Christensen’s urea agar (Becton Dickinson and other sources) is inoculated and incubated aerobically at 35°C for up to 7 days. Development of a pink color indicates a positive reaction. An alternative format is to dispense Christensen’s medium without agar into a microtiter tray well and, after inoculation, overlay it with mineral oil prior to incubation.

VP Test The VP test can be performed as described above for the identification of beta-hemolytic streptococci. A standard method for performing the VP test, requiring extended incubation, is described in chapter 19.

Strains may be isolated as single colonies on sucrosecontaining agar. The two most commonly used media are (i) mitis-salivarius agar containing 0.001% (wt/vol) potassium tellurite (Becton Dickinson) and (ii) tryptone-yeast-cystine agar (Lab M, Bury, United Kingdom). Incubation may require up to 5 days at 37°C under anaerobic incubation conditions.

TYPING SYSTEMS In the majority of cases, typing of streptococci has no immediate clinical or therapeutic consequences. It is most often performed by reference laboratories for the purposes of epidemiologic studies and the evaluation of vaccine efficacy. Although classical antibody-dependent typing systems for capsular serotypes and surface proteins have been used for years, molecular methods are increasingly applied, since they do not require special techniques or the maintenance of rarely used reagents such as a large antibody panel. Another advantage is the independence of DNA sequences from culture conditions and gene expression. For the differentiation of distinct clones, pulsed-field gel electrophoresis (PFGE) and multilocus sequence typing (MLST) systems have been established for many streptococcal species (121). S. pneumoniae comprises more than 90 antigenically distinct capsular serotypes that can be detected by the Neufeld test (Quellung reaction), which is still regarded as the gold standard for epidemiologic studies. Pure cultures of pneumococci are grown on a freshly prepared 5% sheep blood agar plate or a 10% horse blood agar plate at 35°C to 37°C and 5% CO2 for 18 to 24 h. A small amount of bacterial growth (less than a 10-μl loopful) is resuspended in a droplet of phosphate-buffered or physiological saline (McFarland standard of ∼0.5). A few microliters of the saline suspension are mixed with an equal amount of specific pneumococcal rabbit antisera on a glass slide. The specimen is subsequently evaluated for capsular swelling (a clear area surrounding the bacterial cells) by phase-contrast microscopy (oil immersion; magnification, ×1,000) (122). Following the same principle, commercially available kits (Pneumotest Statens Serum Institut, Copenhagen, Denmark) allow rapid testing of S. pneumoniae serotypes with pooled antisera by a checkerboard method. A rapid antigen detection test using pooled antisera coupled to latex beads (Statens Serum Institut) has been developed (123). Due to strain discrepancies, confirmation by the Neufeld Quellung reaction is recommended. In addition, PCR methods for pneumococcal serotype determination have been developed during the last years (124– 126). For the distinction of single clones, PFGE (127) or

396 n

BACTERIOLOGY

MLST typing schemes (128) have been used in pneumococcal investigations. Ten different antibody-defined capsular polysaccharides have been described for S. agalactiae (Ia, Ib, and II to IX). The percentage of nontypeable strains can be minimized by optimization of capsular expression (129). In addition to antibody detection of capsular serotypes, PCR- and DNA sequencing-based techniques allow the detection of capsular serotypes (130, 131). Individual clones of S. agalactiae have been detected either by MLST (132) or by PFGE (133). Conventional typing of S. pyogenes is based upon the antigenic specificity of the surface-expressed T and M proteins (134). The trypsin-resistant T protein is part of the pilus structures (135). The T type can be identified by agglutination with commercially available serologic assays utilizing approximately 20 accepted anti-T sera. M proteins are major antiphagocytic virulence factors of S. pyogenes (136). N-terminal sequence variation in genes encoding these highly protective antigens is the basis of the S. pyogenes precipitation typing system. At present, 83 M serotypes are unequivocally validated and internationally recognized to be serologically unique and are designated M1 to M93 in the Lancefield classification (137). M serotypes that are not included are from non-S. pyogenes organisms or correspond to an existing M serotype. A molecular typing system is based on the nucleotide sequences encoding the amino termini of M proteins. The emm gene sequences encode M proteins and have been correlated with Lancefield M serotypes. This methodology allows assignment to a validated M protein gene sequence (emm1 through emm124) and the identification of new emm sequence types and has evolved into the gold standard of S. pyogenes typing (137). A large database of approximately 350 emm gene sequences from strains originally used for Lancefield serotyping and including emm sequences from beta-hemolytic streptococci of groups C, G, and L is maintained at the CDC (http://www.cdc.gov/ncidod/biotech/ strep/strepblast.htm). An MLST scheme is also available for S. pyogenes. Population genetic studies demonstrated stable associations between emm type and MLST among isolates obtained decades apart and/or from different continents (138). In outbreak situations that include S. pyogenes, restriction enzyme-mediated digestion of emm amplicons is a valuable tool for rapid identification of isolates containing similar emm genes (139). For clusters of isolates sharing the same emm type, PFGE profiles may be helpful for distinguishing similar strains (140).

SEROLOGIC TESTS Determination of streptococcal antibodies is indicated for the diagnosis of poststreptococcal disease, such as acute rheumatic fever or glomerulonephritis (141). A 4-fold rise in antibody titer is regarded as definitive proof of an antecedent streptococcal infection. Multiple variables, including site of infection, time since the onset of infection, age, background prevalence of streptococcal infections (142), antimicrobial therapy, and other comorbidities, influence antibody levels. The most widely used antibodies are anti-streptolysin O (anti-ASO) and anti-DNase B. Antibodies against ASO reach a maximum at 3 to 6 weeks after infection. While ASO responses following streptococcal upper respiratory tract infections are usually elevated, pyoderma caused by S. pyogenes does not elicit a strong ASO response. Streptococcus dysgalactiae subsp. equi-

similis can also produce streptolysin O, and thus elevated ASO titers are not specific for S. pyogenes infections. Among the four streptococcal DNases produced, the host response is most consistent against DNase B. Anti-DNase B titers may not reach maximum titers for 6 to 8 weeks but remain elevated longer than ASO titers and are more reliable than ASO for the confirmation of a preceding streptococcal skin infection. Moreover, since only 80 to 85% of patients with rheumatic fever have elevated ASO titers, additional anti-DNase B titers may be helpful. Due to frequent exposure to S. pyogenes, ASO and antiDNase B titers are higher in children in the United States from 2 to 12 years of age. Geometric mean values are 89 Todd Units for ASO and 112 Units for anti-DNase B, while the upper limits of normal values are 240 Todd Units (ASO) and 640 Units (anti DNAse B) (143). Prompt antibiotic therapy of streptococcal infections can reduce the titer but does not abolish antibody production. Streptococcal carriers do not experience a rise in streptococcal antibody titers. The hemagglutination-based Streptozyme test (CarterWallace, Inc., Cranbury, NJ) was developed to detect antibodies against multiple extracellular streptococcal products. However, variabilities in test standardization and inconsistent specificities have been reported (144). Antibody detection against other S. pyogenes proteins (hyluronidase, streptokinase, and NAD glycohydrolase [NADase]) are technically difficult to perform and not commercially available.

ANTIMICROBIAL SUSCEPTIBILITIES Beta-Hemolytic Streptococci Penicillin remains the drug of choice for the empirical treatment of streptococcal infections due to S. pyogenes, because in contrast to S. pneumoniae and other alpha-hemolytic streptococci, S. pyogenes remains uniformly susceptible to penicillin. Reports about reduced penicillin susceptibility in strains of S. pyogenes have not been confirmed by reference laboratories. This is, however, no longer true for S. agalactiae. The emergence of diminished susceptibility to penicillin G caused by a mutation of the penicillin binding proteins (PBPs) Pbp2x in isolated strains in Asia and the United States has been reported (145, 146). Due to suspected or confirmed penicillin allergies in more than 10% of patients, macrolides are often given as an alternative treatment. Macrolide resistance rates among isolates of S. pyogenes and S. agalactiae have been increasing in North America as well as in Europe (147). Resistance rates correlate with the use of macrolides in clinical practice, and geographic differences in resistance rates are often due to differences in macrolide use. In the United States, macrolide resistance among S. agalactiae strains rose from 12 to 38% from 1990 to 2006 (148, 149) but has substantially declined to about 5% in one recent investigation (150). While isolates with a reduced susceptibility to glycopeptides have not been found in S. pyogenes, a very recent publication reports two independent S. agalactiae isolates from invasive infections, harboring vanG resistance genes (151). Due to the uniform susceptibility of S. pyogenes to penicillin, resistance testing for penicillins or other β-lactams approved for treatment of S. pyogenes and S. agalactiae is not necessary for clinical purposes. So far, the rare isolates of S. agalactiae with reduced susceptibility to penicillin have not resulted in a change of this recommendation, which may of course change if increasing numbers of such strains are encountered. Susceptibility testing for macrolides should be performed using erythromycin, since resistance and susceptibility of azithromycin, clarithro-

22. Streptococcus

mycin, and dirithromycin can be predicted by testing erythromycin. As specified in the CDC guidelines (35), S. agalactiae isolates from pregnant women with severe penicillin allergy should be tested for erythromycin and clindamycin resistance. Testing should include constitutive as well as inducible clindamycin resistance, preferentially with the double-disk diffusion method (D-zone test). In accordance with changes in the most recent CLSI recommendations, however, only results for clindamycin should be reported.

S. pneumoniae and Viridans Group Streptococci In view of the development of penicillin resistance in S. pneumoniae and other alpha-hemolytic streptococci, penicillin can no longer be recommended as the empirical treatment of choice in many countries. Penicillin is considered a preferred antimicrobial agent only for S. pneumoniae and other alpha-hemolytic streptococci with demonstrated susceptibilities to penicillin. Penicillin resistance in S. pneumoniae is caused by altered PBPs. Approximately 25% of S. pneumoniae isolates from the United States were not fully susceptible to penicillin in 2007 (http://www.cdc.gov/abcs/ reports-findings/surv-reports.html). But the changes of S. pneumoniae breakpoints in nonmeningeal isolates (≤2 μg/ ml, susceptible [S]; 4 to 8 μg/ml, intermediate [I]; ≥8 μg/ml, resistant [R]) for penicillin in CLSI definitions caused this value to drop to less than 10% (152). Currently, 5.9% of strains are reported as penicillin resistant (http://www.cdc .gov/abcs/reports-findings/survreports/spneu12.pdf). Susceptibility to penicillin can be determined by a disk diffusion test with 1 μg of oxacillin. According to the current CLSI guidelines, in all cases where oxacillin zone sizes (≤19 mm) indicate a reduced penicillin susceptibility, MIC determinations for penicillin should be performed. For susceptibility testing of all other β-lactams in S. pneumoniae, MIC determinations are recommended. S. pneumoniae infections should be treated according to current guidelines (153). Depending on the clinical situation, treatment options include penicillin, extended-spectrum cephalosporins, macrolides, fluoroquinolones, and vancomycin. In addition, more than one-third of blood culture isolates of the viridans group streptococci collected in the late 1990s in the United States were not susceptible to penicillin (154). Elevated percentages of penicillin-resistant strains can be found among S. mitis and S. salivarius isolates. S. pneumoniae was uniformly susceptible to macrolides until the late 1980s in the United States, but macrolide resistance rates as high as 29% for S. pneumoniae strains have been reported in more-recent investigations (41, 155). The increased use of fluoroquinolones to treat S. pneumoniae infections has been accompanied by a rise in fluoroquinolone-resistant S. pneumoniae strains. Resistance occurs in a stepwise fashion and is due to mutations in DNA topoisomerase IV (ParC) and/or a subunit of DNA gyrase (GyrA). While the overall prevalence of fluoroquinolone resistance remains below 1% according to the most recent Active Bacterial Core Surveillance (ABCs) data (http:// www.cdc.gov/abcs/reports-findings/survreports/spneu12.pdf), the increase in resistant strains during recent years emphasizes the need for close monitoring. Clinical failures of levofloxacin therapy due to resistance have been reported (156). Vancomycin-resistant S. pneumoniae isolates have not been described. However, the isolation of a vancomycinresistant S. bovis isolate has been reported (157).

EVALUATION, INTERPRETATION, AND REPORTING OF RESULTS Streptococci from the pyogenic group are important human pathogens. Timely identification of these species in clinical

n

397

specimens is therefore crucial to treat infections adequately and to reduce transmission. Tonsillopharyngitis caused by S. pyogenes remains a substantial health problem in childhood and adolescence. Diagnosis by either rapid antigen tests, NAATs, or bacteriological culture minimize unjustified antibiotic treatment of viral pharyngitis. Serologic tests are usually applied in cases of suspected poststreptococcal sequelae. Proper identification and reporting should, however, not be limited to S. pyogenes, since S. dysgalactiae subsp. equisimilis (human group C and G streptococci) has been documented as an agent of pharyngitis, including cases complicated by nonsuppurative sequelae. In this context, it is important to correctly differentiate these pathogens from the small-colony-forming beta-hemolytic species of the S. anginosus group that make up part of the oropharyngeal microbiota. While invasive neonatal S. agalactiae infections are declining due to improved prenatal screening and peripartal antibiotic prophylaxis, increased detection of S. agalactiae from adult patients has been reported (158). Thorough identification and reporting of this organism should therefore not be confined to screening swabs during pregnancy or in newborns. Despite the fact that S. pneumoniae is often found as a colonizer in respiratory samples, it should always be clearly distinguished from viridans group streptococci and reported. Cultural methods remain the mainstay in pneumococcal pneumonia and sepsis as well as meningitis. To enable the initiation of adequate antibiotic treatment, resistance testing should be performed for all isolates. Special care should be taken to ensure the reporting of the correct β-lactam breakpoints for non-CSF and CSF S. pneumoniae isolates. If antibiotic treatment was started before microbiological samples were obtained, the urinary antigen test or nucleic acid detection techniques may help to properly identify the causative agent, especially in invasive infections. The correct identification of viridans group streptococci and the distinction of strains causing infections from isolates of the physiological microbiota remains a major challenge. Identification to the group or species level should be confined to strains causing abscesses, endocarditis, and serious infections in neutropenic patients. Many S. mitis isolates are no longer penicillin susceptible, and special attention has to be paid to susceptibility testing. Due to the association of S. gallolyticus subsp. gallolyticus with malignancies of the gastrointestinal tract, and in view of the taxonomic changes within the S. bovis group, reports of novel species designations should include the information that the species belongs to the S. bovis group (159).

REFERENCES 1. Facklam R. 2002. What happened to the streptococci: overview of taxonomic and nomenclature changes. Clin Microbiol Rev 15:613–630. 2. Kohler W. 2007. The present state of species within the genera Streptococcus and Enterococcus. Int J Med Microbiol 297:133–150. 3. Brandt CM, Haase G, Schnitzler N, Zbinden R, Lutticken R. 1999. Characterization of blood culture isolates of Streptococcus dysgalactiae subsp. equisimilis possessing Lancefield’s group A antigen. J Clin Microbiol 37:4194– 4197. 4. Kawamura Y, Hou XG, Sultana F, Miura H, Ezaki T. 1995. Determination of 16S rRNA sequences of Streptococcus mitis and Streptococcus gordonii and phylogenetic relationships among members of the genus Streptococcus. Int J Syst Bacteriol 45:406–408. 5. Kilian M, Poulsen K, Blomqvist T, Havarstein LS, BekThomsen M, Tettelin H, Sorensen UB. 2008. Evolution

398 n

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

20.

BACTERIOLOGY

of Streptococcus pneumoniae and its close commensal relatives. PLoS One 3:e2683. Arbique JC, Poyart C, Trieu-Cuot P, Quesne G, Carvalho Mda G, Steigerwalt AG, Morey RE, Jackson D, Davidson RJ, Facklam RR. 2004. Accuracy of phenotypic and genotypic testing for identification of Streptococcus pneumoniae and description of Streptococcus pseudopneumoniae sp. nov. J Clin Microbiol 42:4686–4696. Keith ER, Podmore RG, Anderson TP, Murdoch DR. 2006. Characteristics of Streptococcus pseudopneumoniae isolated from purulent sputum samples. J Clin Microbiol 44:923–927. Kawamura Y, Hou XG, Todome Y, Sultana F, Hirose K, Shu SE, Ezaki T, Ohkuni H. 1998. Streptococcus peroris sp. nov. and Streptococcus infantis sp. nov., new members of the Streptococcus mitis group, isolated from human clinical specimens. Int J Syst Bacteriol 48(Part 3):921–927. Willcox MD, Zhu H, Knox KW. 2001. Streptococcus australis sp. nov., a novel oral streptococcus. Int J Syst Evol Microbiol 51:1277–1281. Tong H, Gao X, Dong X. 2003. Streptococcus oligofermentans sp. nov., a novel oral isolate from caries-free humans. Int J Syst Evol Microbiol 53:1101–1104. Glazunova OO, Raoult D, Roux V. 2006. Streptococcus massiliensis sp. nov., isolated from a patient blood culture. Int J Syst Evol Microbiol 56:1127–1131. Zbinden A, Mueller NJ, Tarr PE, Sproer C, Keller PM, Bloemberg GV. 2012. Streptococcus tigurinus sp. nov., isolated from blood of patients with endocarditis, meningitis and spondylodiscitis. Int J Syst Evol Microbiol 62:2941– 2945. Martin V, Manes-Lazaro R, Rodriguez JM, MaldonadoBarragan A. 2011. Streptococcus lactarius sp. nov., isolated from breast milk of healthy women. Int J Syst Evol Microbiol 61:1048–1052. Kilian M, Mikkelsen L, Henrichsen J. 1989. Taxonomic study of viridans streptococci: description of Streptococcus gordonii sp. nov. and emended descriptions of Streptococcus sanguis (White and Niven 1946) Streptococcus oralis (Bridge and Sneath 1982) and Streptococcus mitis (Andrews and Horder 1906). Int J Syst Bacteriol 39:471–484. Whiley RA, Beighton D. 1991. Emended descriptions and recognition of Streptococcus constellatus, Streptococcus intermedius, and Streptococcus anginosus as distinct species. Int J Syst Bacteriol 41:1–5. Jensen A, Hoshino T, Kilian M. 2013. Taxonomy of the Anginosus group of the genus Streptococcus and description of Streptococcus anginosus subsp. whileyi subsp. nov. and Streptococcus constellatus subsp. viborgensis subsp. nov. Int J Syst Evol Microbiol 63:2506–2519. Schlegel L, Grimont F, Ageron E, Grimont PA, Bouvet A. 2003. Reappraisal of the taxonomy of the Streptococcus bovis/Streptococcus equinus complex and related species: description of Streptococcus gallolyticus subsp. gallolyticus subsp. nov., S. gallolyticus subsp. macedonicus subsp. nov. and S. gallolyticus subsp. pasteurianus subsp. nov. Int J Syst Evol Microbiol 53:631–645. Devriese LA, Vandamme P, Pot B, Vanrobaeys M, Kersters K, Haesebrouck F. 1998. Differentiation between Streptococcus gallolyticus strains of human clinical and veterinary origins and Streptococcus bovis strains from the intestinal tracts of ruminants. J Clin Microbiol 36:3520–3523. Schlegel L, Grimont F, Collins MD, Regnault B, Grimont PA, Bouvet A. 2000. Streptococcus infantarius sp. nov., Streptococcus infantarius subsp. infantarius subsp. nov. and Streptococcus infantarius subsp. coli subsp. nov., isolated from humans and food. Int J Syst Evol Microbiol 50 Pt 4:1425– 1434. Rurangirwa FR, Teitzel CA, Cui J, French DM, McDonough PL, Besser T. 2000. Streptococcus didelphis sp. nov.,

21.

22.

23.

24.

25.

26.

27.

28.

29. 30. 31.

32. 33.

34.

35.

36.

37.

a streptococcus with marked catalase activity isolated from opossums (Didelphis virginiana) with suppurative dermatitis and liver fibrosis. Int J Syst Evol Microbiol 50(Part 2):759–765. Greenberg D, Broides A, Blancovich I, Peled N, GivonLavi N, Dagan R. 2004. Relative importance of nasopharyngeal versus oropharyngeal sampling for isolation of Streptococcus pneumoniae and Haemophilus influenzae from healthy and sick individuals varies with age. J Clin Microbiol 42:4604–4609. Regev-Yochay G, Raz M, Dagan R, Porat N, Shainberg B, Pinco E, Keller N, Rubinstein E. 2004. Nasopharyngeal carriage of Streptococcus pneumoniae by adults and children in community and family settings. Clin Infect Dis 38:632– 639. Hendley JO, Sande MA, Stewart PM, Gwaltney JM, Jr. 1975. Spread of Streptococcus pneumoniae in families. I. Carriage rates and distribution of types. J Infect Dis 132:55– 61. Schuchat A, Hilger T, Zell E, Farley MM, Reingold A, Harrison L, Lefkowitz L, Danila R, Stefonek K, Barrett N, Morse D, Pinner R. 2001. Active bacterial core surveillance of the emerging infections program network. Emerg Infect Dis 7:92–99. Fulde M, Valentin-Weigand P. 2013. Epidemiology and pathogenicity of zoonotic streptococci. Curr Top Microbiol Immunol 368:49–81. Vandamme P, Pot B, Falsen E, Kersters K, Devriese LA. 1996. Taxonomic study of lancefield streptococcal groups C, G, and L (Streptococcus dysgalactiae) and proposal of S. dysgalactiae subsp. equisimilis subsp. nov. Int J Syst Bacteriol 46:774–781. Vieira V, Teixeira L, Zahner V, Momen H, Facklam R, Steigerwalt A, Brenner D, Castro A. 1998. Genetic relationships among the different phenotypes of Streptococcus dysgalactiae strains. Int J Syst Bacteriol 48:1231–1243. Brandt CM, Spellerberg B. 2009. Human infections due to Streptococcus dysgalactiae subspecies equisimilis. Clin Infect Dis 49:766–772. Cunningham MW. 2000. Pathogenesis of group A streptococcal infections. Clin Microbiol Rev 13:470–511. Efstratiou A. 2000. Group A streptococci in the 1990s. J Antimicrob Chemother 45(Suppl):3–12. Swedo SE, Leonard HL, Mittleman BB, Allen AJ, Rapoport JL, Dow SP, Kanter ME, Chapman F, Zabriskie J. 1997. Identification of children with pediatric autoimmune neuropsychiatric disorders associated with streptococcal infections by a marker associated with rheumatic fever. Am J Psychiatry 154:110–112. Stevens DL. 2001. Invasive streptococcal infections. J Infect Chemother 7:69–80. Musser JM, Gray BM, Schlievert PM, Pichichero ME. 1992. Streptococcus pyogenes pharyngitis: characterization of strains by multilocus enzyme genotype, M and T protein serotype, and pyrogenic exotoxin gene probing. J Clin Microbiol 30:600–603. Davies HD, McGeer A, Schwartz B, Green K, Cann D, Simor AE, Low DE. 1996. Invasive group A streptococcal infections in Ontario, Canada. Ontario Group A Streptococcal Study Group. N Engl J Med 335:547–554. Verani JR, McGee L, Schrag SJ. 2010. Prevention of perinatal group B streptococcal disease—revised guidelines from CDC, 2010. MMWR Recomm Rep 59:1–36. Farley M. 1995. Group B streptococcal infection in older patients. Spectrum of disease and management strategies. Drugs Aging 6:293–300. Barnham M, Thornton TJ, Lange K. 1983. Nephritis caused by Streptococcus zooepidemicus (Lancefield group C). Lancet i:945–948.

22. Streptococcus 38. Haidan A, Talay SR, Rohde M, Sriprakash KS, Currie BJ, Chhatwal GS. 2000. Pharyngeal carriage of group C and group G streptococci and acute rheumatic fever in an Aboriginal population. Lancet 356:1167–1169. 39. CDC. 2008. Invasive pneumococcal disease in children 5 years after conjugate vaccine introduction—eight states, 1998-2005. MMWR Morb Mortal Wkly Rep 57:144–148. 40. Kellner JD, Vanderkooi OG, MacDonald J, Church DL, Tyrrell GJ, Scheifele DW. 2009. Changing epidemiology of invasive pneumococcal disease in Canada, 1998-2007: update from the Calgary-area Streptococcus pneumoniae research (CASPER) study. Clin Infect Dis 49:205–212. 41. Richter SS, Heilmann KP, Dohrn CL, Riahi F, Beekmann SE, Doern GV. 2009. Changing epidemiology of antimicrobial-resistant Streptococcus pneumoniae in the United States, 2004-2005. Clin Infect Dis 48:e23–e33. 42. Dagan R, Patterson S, Juergens C, Greenberg D, GivonLavi N, Porat N, Gurtman A, Gruber WC, Scott DA. 2013. Comparative immunogenicity and efficacy of 13valent and 7-valent pneumococcal conjugate vaccines in reducing nasopharyngeal colonization: a randomized double-blind trial. Clin Infect Dis 57:952–962. 43. Zuccotti G, Mameli C, Daprai L, Garlaschi ML, Dilillo D, Bedogni G, Faccini M, Gramegna M, Torresani E, PneuMi Study G, Ballerini E, Benincaso A, Bonvissuto M, Bricalli D, Brioschi M, Calloni CS, Camiletti MI, Colella G, De Angelis L, Decarlis S, Di Nello F, Dozzi M, Galli E, Gandini V, Giuliani MG, Laviola F, Loda B, Macedoni M, Mazzucchi E, Metta MG, Moscatiello A, Nannini P, Petruzzi M, Picicco D, Picciotti M, Pisanelli S, Porta N, Ramponi G, Redaelli F, Rubini R, Sala N, Saitta V, Scelza G, Tiso RM, Tomasetto M, Torcoletti M, Travaini M, Valentini M, Vessia C. 2014. Serotype distribution and antimicrobial susceptibilities of nasopharyngeal isolates of Streptococcus pneumoniae from healthy children in the 13-valent pneumococcal conjugate vaccine era. Vaccine 32:527–534. 44. Demczuk WH, Martin I, Griffith A, Lefebvre B, McGeer A, Lovgren M, Tyrrell GJ, Desai S, Sherrard L, Adam H, Gilmour M, Zhanel GG, Toronto Bacterial Diseases Network, Canadian Public Health Laboratory Network. 2013. Serotype distribution of invasive Streptococcus pneumoniae in Canada after the introduction of the 13-valent pneumococcal conjugate vaccine, 2010-2012. Can J Microbiol 59:778–788. 45. Richter SS, Heilmann KP, Dohrn CL, Riahi F, Diekema DJ, Doern GV. 2013. Pneumococcal serotypes before and after introduction of conjugate vaccines, United States, 1999-2011. Emerg Infect Dis 19:1074–1083. 46. Bochud PY, Calandra T, Francioli P. 1994. Bacteremia due to viridans streptococci in neutropenic patients: a review. Am J Med 97:256–264. 47. Laupland KB, Ross T, Church DL, Gregson DB. 2006. Population-based surveillance of invasive pyogenic streptococcal infection in a large Canadian region. Clin Microbiol Infect 12:224–230. 48. Parkins MD, Sibley CD, Surette MG, Rabin HR. 2008. The Streptococcus milleri group—an unrecognized cause of disease in cystic fibrosis: a case series and literature review. Pediatr Pulmonol 43:490–497. 49. Ruoff KL, Miller SI, Garner CV, Ferraro MJ, Calderwood SB. 1989. Bacteremia with Streptococcus bovis and Streptococcus salivarius: clinical correlates of more accurate identification of isolates. J Clin Microbiol 27:305–308. 50. Corredoira JC, Alonso MP, Garcia JF, Casariego E, Coira A, Rodriguez A, Pita J, Louzao C, Pombo B, Lopez MJ, Varela J. 2005. Clinical characteristics and significance of Streptococcus salivarius bacteremia and Streptococcus bovis bacteremia: a prospective 16-year study. Eur J Clin Microbiol Infect Dis 24:250–255.

n

399

51. Smith DJ. 2002. Dental caries vaccines: prospects and concerns. Crit Rev Oral Biol Med 13:335–349. 52. Caufield PW, Cutter GR, Dasanayake AP. 1993. Initial acquisition of mutans streptococci by infants: evidence for a discrete window of infectivity. J Dent Res 72:37–45. 53. Beck M, Frodl R, Funke G. 2008. Comprehensive study of strains previously designated Streptococcus bovis consecutively isolated from human blood cultures and emended description of Streptococcus gallolyticus and Streptococcus infantarius subsp. coli. J Clin Microbiol 46:2966– 2972. 54. Gavin PJ, Thomson RB, Jr, Horng SJ, Yogev R. 2003. Neonatal sepsis caused by Streptococcus bovis variant (biotype II/2): report of a case and review. J Clin Microbiol 41:3433–3435. 55. Klein RS, Recco RA, Catalano MT, Edberg SC, Casey JI, Steigbigel NH. 1977. Association of Streptococcus bovis with carcinoma of the colon. N Engl J Med 297:800–802. 56. Facklam R, Elliott J, Pigott N, Franklin AR. 1995. Identification of Streptococcus porcinus from human sources. J Clin Microbiol 33:385–388. 57. Bekal S, Gaudreau C, Laurence RA, Simoneau E, Raynal L. 2006. Streptococcus pseudoporcinus sp. nov., a novel species isolated from the genitourinary tract of women. J Clin Microbiol 44:2584–2586. 58. Stoner KA, Rabe LK, Austin MN, Meyn LA, Hillier SL. 2011. Incidence and epidemiology of Streptococcus pseudoporcinus in the genital tract. J Clin Microbiol 49:883–886. 59. Facklam R, Elliott J, Shewmaker L, Reingold A. 2005. Identification and characterization of sporadic isolates of Streptococcus iniae isolated from humans. J Clin Microbiol 43:933–937. 60. Weinstein MR, Litt M, Kertesz DA, Wyper P, Rose D, Coulter M, McGeer A, Facklam R, Ostach C, Willey BM, Borczyk A, Low DE. 1997. Invasive infections due to a fish pathogen, Streptococcus iniae. S. iniae Study Group. N Engl J Med 337:589–594. 61. Lau SK, Curreem SO, Lin CC, Fung AM, Yuen KY, Woo PC. 2013. Streptococcus hongkongensis sp. nov., isolated from a patient with an infected puncture wound and from a marine flatfish. Int J Syst Evol Microbiol 63:2570– 2576. 62. Martin DR, Stanhope JM, Finch LA. 1977. Delayed culture of group-A streptococci: an evaluation of variables in methods of examining throat swabs. J Med Microbiol 10:249–253. 63. Uhl JR, Adamson SC, Vetter EA, Schleck CD, Harmsen WS, Iverson LK, Santrach PJ, Henry NK, Cockerill FR. 2003. Comparison of LightCycler PCR, rapid antigen immunoassay, and culture for detection of group A streptococci from throat swabs. J Clin Microbiol 41:242–249. 64. Gerber MA, Shulman ST. 2004. Rapid diagnosis of pharyngitis caused by group A streptococci. Clin Microbiol Rev 17:571–580. 65. Bisno AL, Gerber MA, Gwaltney JM, Jr, Kaplan EL, Schwartz RH. 2002. Practice guidelines for the diagnosis and management of group A streptococcal pharyngitis. Infectious Diseases Society of America. Clin Infect Dis 35:113–125. 66. Shulman ST, Bisno AL, Clegg HW, Gerber MA, Kaplan EL, Lee G, Martin JM, Van Beneden C. 2012. Clinical practice guideline for the diagnosis and management of group A streptococcal pharyngitis: 2012 update by the Infectious Diseases Society of America. Clin Infect Dis 55:1279–1282. 67. Chapin KC, Blake P, Wilson CD. 2002. Performance characteristics and utilization of rapid antigen test, DNA probe, and culture for detection of group a streptococci in an acute care clinic. J Clin Microbiol 40:4207–4210.

400 n

BACTERIOLOGY

68. Peterson LR, Thomson RB, Jr. 1999. Use of the clinical microbiology laboratory for the diagnosis and management of infectious diseases related to the oral cavity. Infect Dis Clin North Am 13:775–795. 69. Thinkhamrop J, Limpongsanurak S, Festin MR, Daly S, Schuchat A, Lumbiganon P, Zell E, Chipato T, Win AA, Perilla MJ, Tolosa JE, Whitney CG. 2003. Infections in international pregnancy study: performance of the optical immunoassay test for detection of group B streptococcus. J Clin Microbiol 41:5288–5290. 70. Gutierrez F, Masia M, Rodriguez JC, Ayelo A, Soldan B, Cebrian L, Mirete C, Royo G, Hidalgo AM. 2003. Evaluation of the immunochromatographic Binax NOW assay for detection of Streptococcus pneumoniae urinary antigen in a prospective study of community-acquired pneumonia in Spain. Clin Infect Dis 36:286–292. 71. Roson B, Fernandez-Sabe N, Carratala J, Verdaguer R, Dorca J, Manresa F, Gudiol F. 2004. Contribution of a urinary antigen assay (Binax NOW) to the early diagnosis of pneumococcal pneumonia. Clin Infect Dis 38:222–226. 72. Andreo F, Prat C, Ruiz-Manzano J, Lores L, Blanco S, Cuesta MA, Gimenez M, Dominguez J. 2009. Persistence of Streptococcus pneumoniae urinary antigen excretion after pneumococcal pneumonia. Eur J Clin Microbiol Infect Dis 28:197–201. 73. Dowell SF, Garman RL, Liu G, Levine OS, Yang YH. 2001. Evaluation of Binax NOW, an assay for the detection of pneumococcal antigen in urine samples, performed among pediatric patients. Clin Infect Dis 32:824–825. 74. Mandell LA, Bartlett JG, Dowell SF, File TM, Jr, Musher DM, Whitney C. 2003. Update of practice guidelines for the management of community-acquired pneumonia in immunocompetent adults. Clin Infect Dis 37:1405–1433. 75. Thomas JG. 1994. Routine CSF antigen detection for agents associated with bacterial meningitis: another point of view. Clin Microbiol Newsl 16:89–95. 76. Mein J, Lum G. 1999. CSF bacterial antigen detection tests offer no advantage over Gram’s stain in the diagnosis of bacterial meningitis. Pathology 31:67–69. 77. Moisi JC, Saha SK, Falade AG, Njanpop-Lafourcade BM, Oundo J, Zaidi AK, Afroj S, Bakare RA, Buss JK, Lasi R, Mueller J, Odekanmi AA, Sangare L, Scott JA, Knoll MD, Levine OS, Gessner BD. 2009. Enhanced diagnosis of pneumococcal meningitis with use of the Binax NOW immunochromatographic test of Streptococcus pneumoniae antigen: a multisite study. Clin Infect Dis 48(Suppl 2):S49– S56. 78. Pokorski SJ, Vetter EA, Wollan PC, Cockerill FR, III. 1994. Comparison of Gen-Probe Group A streptococcus Direct Test with culture for diagnosing streptococcal pharyngitis. J Clin Microbiol 32:1440–1443. 79. Anderson NW, Buchan BW, Mayne D, Mortensen JE, Mackey TL, Ledeboer NA. 2013. Multicenter clinical evaluation of the Illumigene group A Streptococcus DNA amplification assay for detection of group A Streptococcus from pharyngeal swabs. J Clin Microbiol 51:1474–1477. 80. Bergeron MG, Ke D, Menard C, Picard FJ, Gagnon M, Bernier M, Ouellette M, Roy PH, Marcoux S, Fraser WD. 2000. Rapid detection of group B streptococci in pregnant women at delivery. N Engl J Med 343:175–179. 81. Davies HD, Miller MA, Faro S, Gregson D, Kehl SC, Jordan JA. 2004. Multicenter study of a rapid molecularbased assay for the diagnosis of group B Streptococcus colonization in pregnant women. Clin Infect Dis 39:1129– 1135. 82. Gavino M, Wang E. 2007. A comparison of a new rapid real-time polymerase chain reaction system to traditional culture in determining group B streptococcus colonization. Am J Obstet Gynecol 197:388 e381–384.

83. Messmer TO, Sampson JS, Stinson A, Wong B, Carlone GM, Facklam RR. 2004. Comparison of four polymerase chain reaction assays for specificity in the identification of Streptococcus pneumoniae. Diagn Microbiol Infect Dis 49:249–254. 84. Carvalho Mda G, Tondella ML, McCaustland K, Weidlich L, McGee L, Mayer LW, Steigerwalt A, Whaley M, Facklam RR, Fields B, Carlone G, Ades EW, Dagan R, Sampson JS. 2007. Evaluation and improvement of realtime PCR assays targeting lytA, ply, and psaA genes for detection of pneumococcal DNA. J Clin Microbiol 45:2460–2466. 85. Albrich WC, Madhi SA, Adrian PV, van Niekerk N, Mareletsi T, Cutland C, Wong M, Khoosal M, Karstaedt A, Zhao P, Deatly A, Sidhu M, Jansen KU, Klugman KP. 2012. Use of a rapid test of pneumococcal colonization density to diagnose pneumococcal pneumonia. Clin Infect Dis 54:601–609. 86. Denys GA, Carey RB. 1992. Identification of Streptococcus pneumoniae with a DNA probe. J Clin Microbiol 30:2725– 2727. 87. Murray PR, Wold AD, Schreck CA, Washington JA, II. 1976. Effects of selective media and atmosphere of incubation on the isolation of group A streptococci. J Clin Microbiol 4:54–56. 88. Bisno AL. 2001. Acute pharyngitis. N Engl J Med 344:205–211. 89. Kellogg JA. 1990. Suitability of throat culture procedures for detection of group A streptococci and as reference standards for evaluation of streptococcal antigen detection kits. J Clin Microbiol 28:165–169. 90. Kurzynski TA, Meise CK. 1979. Evaluation of sulfamethoxazole-trimethoprim blood agar plates for recovery of group A streptococci from throat cultures. J Clin Microbiol 9:189–193. 91. Welch DF, Hensel D, Pickett D, Johnson S. 1991. Comparative evaluation of selective and nonselective culture techniques for isolation of group A beta-hemolytic streptococci. Am J Clin Pathol 95:587–590. 92. Schrag S, Gorwitz R, Fultz-Butts K, Schuchat A. 2002. Prevention of perinatal group B streptococcal disease. Revised guidelines from CDC. MMWR Recomm Rep 51:1– 22. 93. Rosa-Fraile M, Rodriguez-Granger J, Cueto-Lopez M, Sampedro A, Gaye EB, Haro JM, Andreu A. 1999. Use of Granada medium to detect group B streptococcal colonization in pregnant women. J Clin Microbiol 37:2674–2677. 94. Votava M, Tejkalova M, Drabkova M, Unzeitig V, Braveny I. 2001. Use of GBS media for rapid detection of group B streptococci in vaginal and rectal swabs from women in labor. Eur J Clin Microbiol Infect Dis 20:120– 122. 95. Block T, Munson E, Culver A, Vaughan K, Hryciuk JE. 2008. Comparison of carrot broth- and selective ToddHewitt broth-enhanced PCR protocols for real-time detection of Streptococcus agalactiae in prenatal vaginal/anorectal specimens. J Clin Microbiol 46:3615–3620. 96. Richter SS, Heilmann KP, Dohrn CL, Riahi F, Beekmann SE, Doern GV. 2008. Accuracy of phenotypic methods for identification of Streptococcus pneumoniae isolates included in surveillance programs. J Clin Microbiol 46:2184–2188. 97. Schulthess B, Brodner K, Bloemberg GV, Zbinden R, Bottger EC, Hombach M. 2013. Identification of Grampositive cocci by use of matrix-assisted laser desorption ionization-time of flight mass spectrometry: comparison of different preparation methods and implementation of a practical algorithm for routine diagnostics. J Clin Microbiol 51:1834–1840.

22. Streptococcus 98. Malhotra-Kumar S, Wang S, Lammens C, Chapelle S, Goossens H. 2003. Bacitracin-resistant clone of Streptococcus pyogenes isolated from pharyngitis patients in Belgium. J Clin Microbiol 41:5282–5284. 99. Mihaila-Amrouche L, Bouvet A, Loubinoux J. 2004. Clonal spread of emm type 28 isolates of Streptococcus pyogenes that are multiresistant to antibiotics. J Clin Microbiol 42:3844–3846. 100. Pires R, Rolo D, Mato R, Feio de Almeida J, Johansson C, Henriques-Normark B, Morais A, Brito-Avo A, Goncalo-Marques J, Santos-Sanches I. 2009. Resistance to bacitracin in Streptococcus pyogenes from oropharyngeal colonization and noninvasive infections in Portugal was caused by two clones of distinct virulence genotypes. FEMS Microbiol Lett 296:235–240. 101. Kirby R, Ruoff KL. 1995. Cost-effective, clinically relevant method for rapid identification of beta-hemolytic streptococci and enterococci. J Clin Microbiol 33:1154– 1157. 102. Ratner HB, Weeks LS, Stratton CW. 1986. Evaluation of spot CAMP test for identification of group B streptococci. J Clin Microbiol 24:296–297. 103. Hwang MN, Ederer GM. 1975. Rapid hippurate hydrolysis method for presumptive identification of group B streptococci. J Clin Microbiol 1:114–115. 104. Beighton D, Hardie JM, Whiley RA. 1991. A scheme for the identification of viridans streptococci. J Med Microbiol 35:367–372. 105. Kikuchi K, Enari T, Totsuka K, Shimizu K. 1995. Comparison of phenotypic characteristics, DNA-DNA hybridization results, and results with a commercial rapid biochemical and enzymatic reaction system for identification of viridans group streptococci. J Clin Microbiol 33:1215– 1222. 106. Gorm Jensen T, Bossen Konradsen H, Bruun B. 1999. Evaluation of the Rapid ID 32 Strep system. Clin Microbiol Infect 5:417–423. 107. Haanpera M, Jalava J, Huovinen P, Meurman O, Rantakokko-Jalava K. 2007. Identification of alpha-hemolytic streptococci by pyrosequencing the 16S rRNA gene and by use of VITEK 2. J Clin Microbiol 45:762–770. 108. Teles C, Smith A, Ramage G, Lang S. 2011. Identification of clinically relevant viridans group streptococci by phenotypic and genotypic analysis. Eur J Clin Microbiol Infect Dis 30:243–250. 109. Croxatto A, Prod’hom G, Greub G. 2012. Applications of MALDI-TOF mass spectrometry in clinical diagnostic microbiology. FEMS Microbiol Rev 36:380–407. 110. Dubois D, Segonds C, Prere MF, Marty N, Oswald E. 2013. Identification of clinical Streptococcus pneumoniae isolates among other alpha and nonhemolytic streptococci by use of the Vitek MS matrix-assisted laser desorption ionization–time of flight mass spectrometry system. J Clin Microbiol 51:1861–1867. 111. Branda JA, Markham RP, Garner CD, Rychert JA, Ferraro MJ. 2013. Performance of the Vitek MS v2.0 system in distinguishing Streptococcus pneumoniae from nonpneumococcal species of the Streptococcus mitis group. J Clin Microbiol 51:3079–3082. 112. Wojewoda CM, Sercia L, Navas M, Tuohy M, Wilson D, Hall GS, Procop GW, Richter SS. 2013. Evaluation of the Verigene Gram-positive blood culture nucleic acid test for rapid detection of bacteria and resistance determinants. J Clin Microbiol 51:2072–2076. 113. Altun O, Almuhayawi M, Ullberg M, Ozenci V. 2013. Clinical evaluation of the FilmArray blood culture identification panel in identification of bacteria and yeasts from positive blood culture bottles. J Clin Microbiol 51:4130– 4136. 114. Poyart C, Quesne G, Coulon S, Berche P, Trieu-Cuot P. 1998. Identification of streptococci to species level by

n

401

sequencing the gene encoding the manganese-dependent superoxide dismutase. J Clin Microbiol 36:41–47. 115. Whiley RA, Hall LM, Hardie JM, Beighton D. 1999. A study of small-colony, beta-haemolytic, Lancefield group C streptococci within the anginosus group: description of Streptococcus constellatus subsp. pharyngis subsp. nov., associated with the human throat and pharyngitis. Int J Syst Bacteriol 49(Part 4):1443–1449. 116. Summanen PH, Rowlinson MC, Wooton J, Finegold SM. 2009. Evaluation of genotypic and phenotypic methods for differentiation of the members of the Anginosus group streptococci. Eur J Clin Microbiol Infect Dis. 28:1123–1128. 117. Whiley RA, Fraser H, Hardie JM, Beighton D. 1990. Phenotypic differentiation of Streptococcus intermedius, Streptococcus constellatus, and Streptococcus anginosus strains within the “Streptococcus milleri group”. J Clin Microbiol 28:1497–1501. 118. Chuard C, Reller LB. 1998. Bile-esculin test for presumptive identification of enterococci and streptococci: effects of bile concentration, inoculation technique, and incubation time. J Clin Microbiol 36:1135–1136. 119. West PW, Foster HA, Electricwala Q, Alex A. 1996. Comparison of five methods for the determination of arginine hydrolysis by viridans streptococci. J Med Microbiol 45:501–504. 120. Homer KA, Denbow L, Whiley RA, Beighton D. 1993. Chondroitin sulfate depolymerase and hyaluronidase activities of viridans streptococci determined by a sensitive spectrophotometric assay. J Clin Microbiol 31:1648–1651. 121. Maiden MC. 2006. Multilocus sequence typing of bacteria. Annu Rev Microbiol 60:561–588. 122. Austrian R. 1976. The quellung reaction, a neglected microbiologic technique. Mt Sinai J Med 43:699–709. 123. Slotved HC, Kaltoft M, Skovsted IC, Kerrn MB, Espersen F. 2004. Simple, rapid latex agglutination test for serotyping of pneumococci (Pneumotest-Latex). J Clin Microbiol 42:2518–2522. 124. Massire C, Gertz RE, Jr, Svoboda P, Levert K, Reed MS, Pohl J, Kreft R, Li F, White N, Ranken R, Blyn LB, Ecker DJ, Sampath R, Beall B. 2012. Concurrent serotyping and genotyping of pneumococci by use of PCR and electrospray ionization mass spectrometry. J Clin Microbiol 50:2018–2025. 125. Kong F, Brown M, Sabananthan A, Zeng X, Gilbert GL. 2006. Multiplex PCR-based reverse line blot hybridization assay to identify 23 Streptococcus pneumoniae polysaccharide vaccine serotypes. J Clin Microbiol 44:1887–1891. 126. da Gloria Carvalho M, Pimenta FC, Jackson D, Roundtree A, Ahmad Y, Millar EV, O’Brien KL, Whitney CG, Cohen AL, Beall BW. 2010. Revisiting pneumococcal carriage by use of broth enrichment and PCR techniques for enhanced detection of carriage and serotypes. J Clin Microbiol 48:1611–1618. 127. Lefevre JC, Faucon G, Sicard AM, Gasc AM. 1993. DNA fingerprinting of Streptococcus pneumoniae strains by pulsed-field gel electrophoresis. J Clin Microbiol 31:2724– 2728. 128. Enright MC, Spratt BG. 1999. Multilocus sequence typing. Trends Microbiol 7:482–487. 129. Benson JA, Flores AE, Baker CJ, Hillier SL, Ferrieri P. 2002. Improved methods for typing nontypeable isolates of group B streptococci. Int J Med Microbiol 292:37– 42. 130. Kong F, Gowan S, Martin D, James G, Gilbert GL. 2002. Serotype identification of group B streptococci by PCR and sequencing. J Clin Microbiol 40:216–226. 131. Poyart C, Tazi A, Reglier-Poupet H, Billoet A, Tavares N, Raymond J, Trieu-Cuot P. 2007. Multiplex PCR assay for rapid and accurate capsular typing of group B streptococci. J Clin Microbiol 45:1985–1988.

402 n

BACTERIOLOGY

132. Jones N, Bohnsack JF, Takahashi S, Oliver KA, Chan MS, Kunst F, Glaser P, Rusniok C, Crook DW, Harding RM, Bisharat N, Spratt BG. 2003. Multilocus sequence typing system for group B streptococcus. J Clin Microbiol 41:2530–2536. 133. Benson JA, Ferrieri P. 2001. Rapid pulsed-field gel electrophoresis method for group B streptococcus isolates. J Clin Microbiol 39:3006–3008. 134. Johnson DR, Kaplan EL. 1993. A review of the correlation of T-agglutination patterns and M-protein typing and opacity factor production in the identification of group A streptococci. J Med Microbiol 38:311–315. 135. Mora M, Bensi G, Capo S, Falugi F, Zingaretti C, Manetti AG, Maggi T, Taddei AR, Grandi G, Telford JL. 2005. Group A Streptococcus produce pilus-like structures containing protective antigens and Lancefield T antigens. Proc Natl Acad Sci USA 102:15641–15646. 136. Fischetti VA. 1989. Streptococcal M protein: molecular design and biological behavior. Clin Microbiol Rev 2:285– 314. 137. Facklam RF, Martin DR, Lovgren M, Johnson DR, Efstratiou A, Thompson TA, Gowan S, Kriz P, Tyrrell GJ, Kaplan E, Beall B. 2002. Extension of the Lancefield classification for group A streptococci by addition of 22 new M protein gene sequence types from clinical isolates: emm103 to emm124. Clin Infect Dis 34:28–38. 138. Enright MC, Spratt BG, Kalia A, Cross JH, Bessen DE. 2001. Multilocus sequence typing of Streptococcus pyogenes and the relationships between emm type and clone. Infect Immun 69:2416–2427. 139. Beall B, Facklam RR, Elliott JA, Franklin AR, Hoenes T, Jackson D, Laclaire L, Thompson T, Viswanathan R. 1998. Streptococcal emm types associated with Tagglutination types and the use of conserved emm gene restriction fragment patterns for subtyping group A streptococci. J Med Microbiol 47:893–898. 140. Musser JM, Kapur V, Szeto J, Pan X, Swanson DS, Martin DR. 1995. Genetic diversity and relationships among Streptococcus pyogenes strains expressing serotype M1 protein: recent intercontinental spread of a subclone causing episodes of invasive disease. Infect Immun 63:994– 1003. 141. Shet A, Kaplan EL. 2002. Clinical use and interpretation of group A streptococcal antibody tests: a practical approach for the pediatrician or primary care physician. Pediatr Infect Dis J 21:420–426; quiz 427–430. 142. Ayoub EM, Nelson B, Shulman ST, Barrett DJ, Campbell JD, Armstrong G, Lovejoy J, Angoff GH, Rockenmacher S. 2003. Group A streptococcal antibodies in subjects with or without rheumatic fever in areas with high or low incidences of rheumatic fever. Clin Diagn Lab Immunol 10:886–890. 143. Kaplan EL, Rothermel CD, Johnson DR. 1998. Antistreptolysin O and anti-deoxyribonuclease B titers: normal values for children ages 2 to 12 in the United States. Pediatrics 101:86–88. 144. Gerber MA, Wright LL, Randolph MF. 1987. Streptozyme test for antibodies to group A streptococcal antigens. Pediatr Infect Dis J 6:36–40. 145. Dahesh S, Hensler ME, Van Sorge NM, Gertz RE, Jr, Schrag S, Nizet V, Beall BW. 2008. Point mutation in the group B streptococcal pbp2x gene conferring decreased susceptibility to beta-lactam antibiotics. Antimicrob Agents Chemother 52:2915–2918. 146. Kimura K, Suzuki S, Wachino J, Kurokawa H, Yamane K, Shibata N, Nagano N, Kato H, Shibayama K, Arakawa Y. 2008. First molecular characterization of group B streptococci with reduced penicillin susceptibility. Antimicrob Agents Chemother 52:2890–2897.

147. Desjardins M, Delgaty KL, Ramotar K, Seetaram C, Toye B. 2004. Prevalence and mechanisms of erythromycin resistance in group A and group B Streptococcus: implications for reporting susceptibility results. J Clin Microbiol 42:5620–5623. 148. Murdoch DR, Reller LB. 2001. Antimicrobial susceptibilities of group B streptococci isolated from patients with invasive disease: 10-year perspective. Antimicrob Agents Chemother 45:3623–3624. 149. Gygax SE, Schuyler JA, Kimmel LE, Trama JP, Mordechai E, Adelson ME. 2006. Erythromycin and clindamycin resistance in group B streptococcal clinical isolates. Antimicrob Agents Chemother 50:1875–1877. 150. Villasenor-Sierra A, Katahira E, Jaramillo-Valdivia AN, Barajas-Garcia Mde L, Bryant A, Morfin-Otero R, Marquez-Diaz F, Tinoco JC, Sanchez-Corona J, Stevens DL. 2012. Phenotypes and genotypes of erythromycin-resistant Streptococcus pyogenes strains isolated from invasive and non-invasive infections from Mexico and the USA during 1999-2010. Int J Infect Dis 16:e178–e181. 151. Park C, Nichols M, Schrag SJ. 2014. Two cases of invasive vancomycin-resistant group B streptococcus infection. N Engl J Med 370:885–886. 152. Weinstein MP, Klugman KP, Jones RN. 2009. Rationale for revised penicillin susceptibility breakpoints versus Streptococcus pneumoniae: coping with antimicrobial susceptibility in an era of resistance. Clin Infect Dis 48:1596– 1600. 153. Mandell LA, Wunderink RG, Anzueto A, Bartlett JG, Campbell GD, Dean NC, Dowell SF, File TM, Jr, Musher DM, Niederman MS, Torres A, Whitney CG. 2007. Infectious Diseases Society of America/American Thoracic Society consensus guidelines on the management of community-acquired pneumonia in adults. Clin Infect Dis 44 Suppl 2:S27–S72. 154. Doern GV, Ferraro MJ, Brueggemann AB, Ruoff KL. 1996. Emergence of high rates of antimicrobial resistance among viridans group streptococci in the United States. Antimicrob Agents Chemother 40:891–894. 155. Thornsberry C, Sahm DF, Kelly LJ, Critchley IA, Jones ME, Evangelista AT, Karlowsky JA. 2002. Regional trends in antimicrobial resistance among clinical isolates of Streptococcus pneumoniae, Haemophilus influenzae, and Moraxella catarrhalis in the United States: results from the TRUST Surveillance Program, 1999-2000. Clin Infect Dis 34(Suppl 1):S4–S16. 156. Davidson R, Cavalcanti R, Brunton JL, Bast DJ, de Azavedo JC, Kibsey P, Fleming C, Low DE. 2002. Resistance to levofloxacin and failure of treatment of pneumococcal pneumonia. N Engl J Med 346:747–750. 157. Poyart C, Pierre C, Quesne G, Pron B, Berche P, TrieuCuot P. 1997. Emergence of vancomycin resistance in the genus Streptococcus: characterization of a vanB transferable determinant in Streptococcus bovis. Antimicrob Agents Chemother 41:24–29. 158. Skoff TH, Farley MM, Petit S, Craig AS, Schaffner W, Gershman K, Harrison LH, Lynfield R, Mohle-Boetani J, Zansky S, Albanese BA, Stefonek K, Zell ER, Jackson D, Thompson T, Schrag SJ. 2009. Increasing burden of invasive group B streptococcal disease in nonpregnant adults, 1990-2007. Clin Infect Dis 49:85–92. 159. van’t Wout JW, Bijlmer HA. 2005. Bacteremia due to Streptococcus gallolyticus, or the perils of revised nomenclature in bacteriology. Clin Infect Dis 40:1070–1071. 160. Facklam RR, Washington II JA. 1991. Streptococcus and related catalase-negative gram-postiive cocci, p 238–257. In Balows A, Hausler WA, Jr, Herrmann KL, Isenberg HD, Shadomy HJ (ed), Manual of Clinical Microbiology, 5th ed. American Society for Microbiology, Washington, DC.

Enterococcus LÚCIA MARTINS TEIXEIRA, MARIA DA GLÓRIA SIQUEIRA CARVALHO, RICHARD R. FACKLAM, AND PATRICIA LYNN SHEWMAKER

23 16S rRNA gene is a practical and powerful tool in aiding the identification of enterococcal species, and it has been performed for all recognized species of Enterococcus. Figure 1 shows the phylogenetic relationships among the species of Enterococcus based on the analysis of 16S rRNA gene sequences, which are available from the GenBank database. Several other molecular methods, mostly nucleic acid-based assays, have been used as additional tools to assess the phylogenetic relationships among enterococcal species and to formulate the description of new species, but their use is still limited (for further details, see references 1 and 4 and “Identification” below).

TAXONOMY The genus Enterococcus includes microorganisms that have a historical connection with the genus Streptococcus, and their initial documentation is related to the “streptococci of fecal origin” or “enterococci” (see reference 1 for a brief historical overview). They were initially considered a distinct category within the genus Streptococcus, distinguished by their higher resistance to chemical and physical agents and accommodating most of the serological group D streptococci. After the introduction of molecular methods for studying these microorganisms, however, the enterococci have undergone considerable changes in taxonomy, which started with the splitting of the genus Streptococcus and the recognition of Enterococcus as a separate genus in 1984 (2). Streptococcus faecalis and Streptococcus faecium were the first species to be transferred to the new genus as Enterococcus faecalis (the type species) and Enterococcus faecium, respectively. Subsequently, other earlier streptococcal species and subspecies were transferred and received new denominations as members of the genus Enterococcus (3). The continuous use of molecular approaches has allowed major developments in the classification of the enterococci, resulting in the recognition of 49 enterococcal species as of April 2014 (for further details, see references 1 and 4, as well as the List of Prokaryotic Names with Standing in Nomenclature [http:// www.bacterio.net]). The enterococci belong to the low DNA guanine-pluscytosine (G+C 256 (R)

Teicoplanin

16–512 (R)

0.5–1 (S)

0.5–1 (S)

4–64 (S/R)

0.5 (S)

0.5 (S)

0.5 (S)

64 to >256 (R) 0.5 (S)

Classification (level)

High

Variable

Low

Moderate

Low

Low

Low

Variable

Low

Genotype

vanA

vanBb

vanCb

vanDb

vanE

vanGb

vanL

vanM

vanN

Mobile element

Tn1546

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Unknown

Occurrence of conjugation Location of van genes

+

Tn1547 or Tn1549 to Tn5382 +

−

−

+

−

+

+

Plasmid chromosome

Plasmid chromosome

Chromosome

Chromosome

Unknown

Plasmid

Type of expression

Inducible

Inducible

Constitutive

Constitutive

Inducible

Inducible

Unknown

Constitutive

Gene productc (modified target)

D-Ala-D-Lac

D-Ala-D-Lac

D-Ala-D-Ser

D-Ala-D-Lac

D-Ala-D-Ser

D-Ala-D-Ser

Distribution among species

E. faecalis, E. faecium, E. faecalis, E. faecium, E. avium, E. E. durans, E. casseliflavus, E. gallinarum durans, E. gallinarum, E. hirae, E. mundtii, E. raffinosus, E. thailandicus

E. faecalis

E. faecalis

a

R, resistant; S, susceptible; I, intermediate. Subtypes exist: vanB1 to vanB3; vanC1 to vanC4; vanD1 to vanD5; vanG1 to vanG2. Ala, alanine; Lac, lactate; Ser, serine. d Product deduced but not confirmed. b c

−

E.casseliflavus, E. gallinarum

E. faecium, E. faecalis, E. avium, E. gallinarum, E. raffinosus

Chromosome Chromosome Unknown Unknown

12–16 (I)

D-Ala-D-Serd D-Ala-D-Lacd

D-Ala-D-Serd

E. faecalis

E. faecium

E. faecium

23. Enterococcus n 415

occur rarely among enterococci. Furthermore, the isolation of vancomycin-dependent (154) and vancomycin-heteroresistant (155) enterococcal strains from clinically significant infections, although sporadically reported, may also represent additional serious threats for the treatment and control of enterococcal infections. While in vitro methods for detecting vancomycin resistance are discussed in detail in references 1 and 145 as well as in chapters 71 and 73, some aspects regarding vanCcontaining species (e.g., E. gallinarum and E. casseliflavus) need to be emphasized. Resistance associated with vanC genotypes is not usually detected by disk diffusion, but VanC strains frequently grow on vancomycin agar screen tests. Because of the low clinical significance of the VanC resistance, the implications of susceptibility testing for patient management may be unclear. However, the need to differentiate VanA or VanB strains, as well as strains displaying the other less commonly found types of acquired vancomycin resistance, from VanC strains is quite evident for therapeutic, infection control, and surveillance reasons. Because growth on vancomycin screening agar fails to help with this important distinction, species identification is necessary. For practical purposes, growth on the vancomycin screening agar test by E. faecalis or E. faecium is likely due to the presence of vanA or vanB. Although rare, the occurrence of the other kinds of vancomycin resistance may also be considered. Additionally, simultaneous occurrence of the vanA gene has been increasingly reported in vanC-carrying species E. casseliflavus and especially E. gallinarum, so that identification of a species that usually harbors only VanC resistance does not completely rule out moderate to high levels of vancomycin resistance (28, 49, 55). In this regard, determining vancomycin MICs is useful, as VanC resistance frequently results in MICs of 32 μg/ml. In such cases, determination of the genetic elements associated with vancomycin resistance has important epidemiological and infection control implications. Also, resistance to other agents such as ampicillin and aminoglycosides is uncommon among VanC isolates. Moreover, the unexpected finding of simultaneous occurrence of the vanA and vanC1 genes in an E. faecalis isolate from ewe bulk tank milk was recently reported (156), illustrating the increasing complexity for precise determination of glycopeptide resistance and its association with the different enterococcal species. Because of limited alternatives, chloramphenicol, erythromycin, tetracycline (or doxycycline or minocycline), and rifampin may be tested for VRE. Selective testing of quinupristin-dalfopristin, linezolid, and daptomycin, based on the site of infection, is recommended when reporting vancomycin-resistant E. faecium isolates (144, 145). Molecular methods (see chapter 77) have been used to detect specific antimicrobial resistance genes and have substantially contributed to the understanding of the spread of acquired resistance among enterococci, especially resistance to vancomycin. However, because of their high specificity, molecular methods do not detect antimicrobial resistance due to mechanisms not targeted by the testing, including emerging resistance mechanisms.

EVALUATION, INTERPRETATION, AND REPORTING OF RESULTS The diversity and species specificity of acquired antimicrobial resistance traits among enterococcal isolates created an additional need for accurate identification at the species level and for in vitro evaluation of susceptibility to antimicro-

bial agents. The significance of a particular enterococcal isolate is a major factor in determining when antimicrobial testing should be done. Once the need to test a particular isolate has been established, selection of the appropriate antimicrobial agents for testing must be considered on the basis of the site of infection. Testing of antimicrobial agents to which enterococci are intrinsically resistant is contraindicated. The drugs that should not be reported include aminoglycosides at standard concentrations, cephalosporins, clindamycin, and trimethoprim-sulfamethoxazole. They may appear active for enterococci in vitro but are not effective clinically, and as such they should not be reported as drugs to which enterococci are susceptible. Updated guidelines (145) for the selection of antimicrobial agents should be followed for routine testing and reporting. The in vitro methods for detecting antimicrobial resistance in enterococcal isolates were reviewed and summarized in reference 1 and are also discussed in chapters 71 and 73. As already mentioned, testing for high-level aminoglycoside resistance as a predictor of synergy should be done with any enterococcal isolate implicated in infections for which combination therapy is indicated, e.g., from systemic infections. Enterococci are also frequently encountered in polymicrobial infections associated with the gastrointestinal tract or superficial wounds of hospitalized patients. Their pathogenic significance in such settings is uncertain, but susceptibility testing is warranted when predominant or heavy growth is observed. Testing of E. faecalis isolates from lower urinary tract infections is optional, as these infections usually respond to therapy with ampicillin or nitrofurantoin. However, many hospital infection control programs require routine testing as part of surveillance programs for VRE. For those instances when testing a urinary tract isolate is appropriate, ciprofloxacin, fosfomycin, levofloxacin, norfloxacin, and tetracycline could be selected, in addition to nitrofurantoin and ampicillin (28, 36, 142, 143, 145). In cases of treatment failure, testing is always warranted. The findings and conclusions in this chapter are those of the authors and do not necessarily represent the views of the CDC. The use of product names in the manuscript does not imply their endorsement by the U.S. Department of Health and Human Services.

REFERENCES 1. Facklam RR, Carvalho MGS, Teixeira LM. 2002. History, taxonomy, biochemical characteristics, and antibiotic susceptibility testing of enterococci, p 1–54. In Gilmore MS, Clewell DB, Courvalin P, Dunny GM, Murray BE, Rice LB (ed), The Enterococci: Pathogenesis, Molecular Biology, and Antibiotic Resistance. ASM Press, Washington, DC. 2. Schleifer KH, Kilpper-Balz R. 1984. Transfer of Streptococcus faecalis and Streptococcus faecium to the genus Enterococcus nom. rev. as Enterococcus faecalis comb. nov. and Enterococcus faecium comb. nov. Int J Syst Bacteriol 34:31–34. 3. Collins MD, Jones D, Farrow JAE, Kilpper-Balz R, Schleifer KH. 1984. Enterococcus avium nom. rev., comb. nov.; E. casseliflavus nom. rev., comb. nov.; E. durans nom. rev., comb. nov.; E. gallinarum comb. nov.; and E. malodoratus sp. nov. Int J Syst Bacteriol 34:220–223. 4. Švec P, Devriese LA. 2009. Genus I. Enterococcus, p 594– 623. In De Vos P, Garrity GM, Jones D, Krieg NR, Ludwig W, Rainey FA, Schleifer KH, Whitman WB (ed), Bergey’s Manual of Systematic Bacteriology, 2nd ed. Springer, New York, NY. 5. Devriese L, Baele M, Butaye P. 2006. The genus Enterococcus: Taxonomy, p 163–174. In Dworkin M, Falkow S, Rosenberg E, Schleifer KH, Stackebrandt E (ed), The

416 n

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

BACTERIOLOGY

Prokaryotes. A Handbook on the Biology of Bacteria, 3rd ed, vol 4. Springer, New York, NY. Ludwig W, Schleifer KH, Whitman WB. 2009. Family IV. Enterococcaceae fam. nov., p 594. In De Vos P, Garrity GM, Jones D, Krieg NR, Ludwig W, Rainey FA, Schleifer KH, Whitman WB (ed), Bergey’s Manual of Systematic Bacteriology, 2nd ed. Springer, New York, NY. Euzéby J. 2010. List of new names and new combinations previously effectively, but not validly, published. Int J Syst Evol Microbiol 60:469–472. Huycke MM. 2002. Physiology of enterococci, p 133–175. In Gilmore MS, Clewell DB, Courvalin P, Dunny GM, Murray BE, Rice LB (ed), The Enterococci: Pathogenesis, Molecular Biology, and Antibiotic Resistance. ASM Press, Washington, DC. Van Horn K, Tóth C, Kariyama R, Mitsuhata R, Kumon H. 2002. Evaluation of 15 motility media and a direct microscopic method for detection of motility in enterococci. J Clin Microbiol 40:2476–2479. Švec P, Devriese LA, Sedlacek I, Baele M, Vancanneyt M, Haesbrouck F, Swings J, Doskar J. 2001. Enterococcus haemoperoxidus sp. nov. and Enterococcus moraviensis sp. nov., isolated from water. Int J Syst Evol Microbiol 51:1567–1574. Švec P, Vancanneyt M, Sedlácek I, Naser SM, Snauwaert C, Lefebvre K, Hoste B, Swings J. 2006. Enterococcus silesiacus sp. nov. and Enterococcus termitis sp. nov. Int J Syst Evol Microbiol 56:577–581. Švec P, Vandamme P, Bryndová H, Holochová P, Kosina M, Maslanová I, Sedlácek I. 2012. Enterococcus plantarum sp. nov., isolated from plants. Int J Syst Evol Microbiol 62:1499–1505. Bourgogne A, Garsin DA, Qin X, Singh KV, Sillanpaa J, Yerrapragada S, Ding Y, Dugan-Rocha S, Buhay C, Shen H, Chen G, Williams G, Muzny D, Maadani A, Fox KA, Gioia J, Chen L, Shang Y, Arias CA, Nallapareddy SR, Zhao M, Prakash VP, Chowdhury S, Jiang H, Gibbs RA, Murray BE, Highlander SK, Weinstock GM. 2008. Large scale variation in Enterococcus faecalis illustrated by the genome analysis of strain OG1RF. Genome Biol 9:R110. Palmer KL, Godfrey P, Griggs A, Kos VN, Zucker J, Desjardins C, Cerqueira G, Gevers D, Walker S, Wortman J, Feldgarden M, Haas B, Birren B, Gilmore MS. 2012. Comparative genomics of enterococci: variation in Enterococcus faecalis, clade structure in E. faecium, and defining characteristics of E. gallinarum and E. casseliflavus. mBio 3:e00318–11. Qin X, Galloway-Peña JR, Sillanpaa J, Roh JH, Nallapareddy SR, Chowdhury S, Bourgogne A, Choudhury T, Muzny DM, Buhay CJ, Ding Y, Dugan-Rocha S, Liu W, Kovar C, Sodergren E, Highlander S, Petrosino JF, Worley KC, Gibbs RA, Weinstock GM, Murray BE. 2012. Complete genome sequence of Enterococcus faecium strain TX16 and comparative genomic analysis of Enterococcus faecium genomes. BMC Microbiol 12:135. Paulsen IT, Banerjei L, Myers GS, Nelson KE, Seshadri R, Read TD, Fouts DE, Eisen JA, Gill SR, Heidelberg JF, Tettelin H, Dodson RJ, Umayam L, Brinkac L, Beanan M, Daugherty S, DeBoy RT, Durkin S, Kolonay J, Madupu R, Nelson W, Vamathevan J, Tran B, Upton J, Hansen T, Shetty J, Khouri H, Utterback T, Radune D, Ketchum KA, Dougherty BA, Fraser CM. 2003. Role of mobile DNA in the evolution of vancomycin-resistant Enterococcus faecalis. Science 299:2071–2074. Facklam RR, Elliott JA. 1995. Identification, classification, and clinical relevance of catalase-negative, grampositive cocci, excluding the streptococci and enterococci. Clin Microbiol Rev 8:479–495. Teixeira LM, Carvalho MGS, Merquior VL, Steigerwalt AG, Brenner DJ, Facklam RR. 1997. Phenotypic and

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29. 30.

31.

32.

33.

34.

35.

36.

37.

genotypic characterization of Vagococcus fluvialis, including strains isolated from human sources. J Clin Microbiol 35:2778–2781. Jackson CR, Fedorka-Cray PJ, Barrett JB. 2004. Use of a genus- and species-specific multiplex PCR for identification of enterococci. J Clin Microbiol 42:3558–3565. Tannock GW, Cook G. 2002. Enterococci as members of the intestinal microflora of humans, p 101–132. In Gilmore MS, Clewell DB, Courvalin P, Dunny GM, Murray BE, Rice LB (ed), The Enterococci: Pathogenesis, Molecular Biology, and Antibiotic Resistance. ASM Press, Washington, DC. Moles L, Gómez M, Heilig H, Bustos G, Fuentes S, de Vos W, Fernández L, Rodríguez JM, Jiménez E. 2013. Bacterial diversity in meconium of preterm neonates and evolution of their fecal microbiota during the first month of life. PLoS One 8:e66986. Zirakzadeh A, Patel R. 2006. Vancomycin-resistant enterococci: colonization, infection, detection, and treatment. Mayo Clin Proc 81:529–536. Ubeda C, Taur Y, Jenq RR, Equinda MJ, Son T, Samstein M, Viale A, Socci ND, van den Brink MR, Kamboj M, Pamer EG. 2010. Vancomycin-resistant Enterococcus domination of intestinal microbiota is enabled by antibiotic treatment in mice and precedes bloodstream invasion in humans. J Clin Invest 120:4332–4341. Arias CA, Murray BE. 2012. The rise of the Enterococcus: beyond vancomycin resistance. Nat Rev Microbiol 10:266– 278. Franz CM, Stiles ME, Schleifer KH, Holzapfel WH. 2003. Enterococci in foods—a conundrum for food safety. Int J Food Microbiol 88:105–122. Byappanahalli MN, Nevers MB, Korajkic A, Staley ZR, Harwood VJ. 2012. Enterococci in the environment. Microbiol Mol Biol Rev 76:685–706. Baldassarri L, Creti R, Montanaro L, Orefici G, Arciola CR. 2005. Pathogenesis of implant infections by enterococci. Int J Artif Organs 28:1101–1109. Malani PN, Kauffman CA, Zervos MJ. 2002. Enterococcal disease, epidemiology, and treatment, p 385–408. In Gilmore MS, Clewell DB, Courvalin P, Dunny GM, Murray BE, Rice LB (ed), The Enterococci: Pathogenesis, Molecular Biology and Antibiotic Resistance. ASM Press, Washington, DC. Murray BE. 1990. The life and times of the Enterococcus. Clin Microbiol Rev 3:46–65. Paganelli FL, Willems RJ, Leavis HL. 2012. Optimizing future treatment of enterococcal infections: attacking the biofilm? Trends Microbiol 20:40–49. Tendolkar PM, Baghdayan AS, Shankar N. 2003. Pathogenic enterococci: new developments in the 21st century. Cell Mol Life Sci 60:2622–2636. Leavis HL, Bonten MJ, Willems RJ. 2006. Identification of high-risk enterococcal clonal complexes: global dispersion and antibiotic resistance. Curr Opin Microbiol 9:454– 460. Top J, Willems R, Blok H, de Regt M, Jalink K, Troelstra A, Goorhuis B, Bonten M. 2007. Ecological replacement of Enterococcus faecalis by multiresistant clonal complex 17 Enterococcus faecium. Clin Microbiol Infect 13:316–319. Willems RJ, Bonten MJ. 2007. Glycopeptide-resistant enterococci: deciphering virulence, resistance and epidemicity. Curr Opin Infect Dis 20:384–390. van Schaik W, Willems RJ. 2010. Genome-based insights into the evolution of enterococci. Clin Microbiol Infect 16:527–532. Huycke MM, Sahm DF, Gilmore MS. 1998. Multipleresistant enterococci: the nature of the problem and an agenda for the future. Emerg Infect Dis 4:239–249. Sava IG, Heikens E, Huebner J. 2010. Pathogenesis and immunity in enterococcal infections. Clin Microbiol Infect 16:533–540.

23. Enterococcus n 38. Shankar N, Baghdayan AS, Gilmore MS. 2002. Modulation of virulence within a pathogenicity island in vancomycin resistant Enterococcus faecalis. Nature 417:746–750. 39. McBride SM, Fischetti VA, Leblanc DJ, Moellering RC, Jr, Gilmore MS. 2007. Genetic diversity among Enterococcus faecalis. PLoS ONE 2:e582. 40. Donelli G, Guaglianone E. 2004. Emerging role of Enterococcus spp in catheter-related infections: biofilm formation and novel mechanisms of antibiotic resistance. J Vasc Access 5:3–9. 41. Zehnder M, Guggenheim B. 2009. The mysterious appearance of enterococci in filled root canals. Int Endod J 42:277–287. 42. Daw K, Baghdayan AS, Awasthi S, Shankar N. 2012. Biofilm and planktonic Enterococcus faecalis elicit different responses from host phagocytes in vitro. FEMS Immunol Med Microbiol 65:270–282. 43. National Nosocomial Infections Surveillance System. 2000. National Nosocomial Infections Surveillance (NNIS) system report, data summary from January 1992– April 2000, issued June 2000. Am J Infect Control 28:429– 448. 44. Karlowsky JA, Jones ME, Draghi DC, Thornsberry C, Sahm DF, Volturo GA. 2004. Prevalence and antimicrobial susceptibilities of bacteria isolated from blood cultures of hospitalized patients in the United States in 2002. Ann Clin Microbiol Antimicrob 10:3–7. 45. Hidron AI, Edwards JR, Patel J, Horan TC, Sievert DM, Pollock DA, Fridkin SK, National Healthcare Safety Network Team and Participating National Healthcare Safety Network Facilities. 2008. Antimicrobial-resistant pathogens associated with healthcare-associated infections: annual summary of data reported to the National Healthcare Safety Network at the Centers for Disease Control and Prevention, 2006–2007. Infect Control Hosp Epidemiol 29:996–1011. 46. Sievert DM, Ricks P, Edwards JR, Schneider A, Patel J, Srinivasan A, Kallen A, Limbago B, Fridkin S, National Healthcare Safety Network (NHSN) Team and Participating NHSN Facilities. 2013. Antimicrobial-resistant pathogens associated with healthcare-associated infections: summary of data reported to the National Healthcare Safety Network at the Centers for Disease Control and Prevention, 2009–2010. Infect Control Hosp Epidemiol 34:1– 14. 47. Facklam RR, Collins MD. 1989. Identification of Enterococcus species isolated from human infections by a conventional test scheme. J Clin Microbiol 27:731–734. 48. Gordon S, Swenson JS, Hill BC, Pigott NE, Facklam RR, Cooksey RC, Thornsberry C, the Enterococcal Study Group, Jarvis WR, Tenover FC. 1992. Antimicrobial susceptibility patterns of common and unusual species of enterococci causing infections in the United States. J Clin Microbiol 30:2373–2378. 49. Buschelman BJ, Bale BJ, Jones RN. 1993. Species identification and determination of high-level aminoglycoside resistance among enterococci. Comparison study of sterile body fluid isolates, 1985–1991. Diagn Microbiol Infect Dis 16:119–922. 50. Iwen PC, Kelly DM, Linder J, Hinrichs SH, Dominguez EA, Rupp ME, Patil KD. 1997. Change in prevalence and antibiotic resistance of Enterococcus species isolated from blood cultures over an 8-year period. Antimicrob Agents Chemother 41:494–495. 51. European Centre for Disease Prevention and Control (ECDC). 2013. Point Prevalence Survey of HealthcareAssociated Infections and Antimicrobial Use in European Acute Care Hospitals 2011–2012. ECDC, Stockholm, Sweden. doi:10.2900/86011. http://www.ecdc.europa.eu/en/publica tions/Publications/Forms/ECDC_DispForm.aspx?ID= 1155.

417

52. Nauschuetz WF, Trevino SB, Harrison LS, Longfield RN, Fletcher L, Wortham WG. 1993. Enterococcus casseliflavus as an agent of nosocomial bloodstream infections. Med Microbiol Lett 2:102–108. 53. Contreras GA, DiazGranados CA, Cortes L, Reyes J, Vanegas S, Panesso D, Rincón S, Díaz L, Prada G, Murray BE, Arias CA. 2008. Nosocomial outbreak of Enteroccocus gallinarum: untaming of rare species of enterococci. J Hosp Infect 70:346–352. 54. Merquior VL, Neves FPG, Ribeiro RL, Duarte RS, Marques EA, Teixeira LM. 2008. Bacteraemia associated with a vancomycin-resistant Enterococcus gallinarum strain harboring both the vanA and vanC1 genes. J Med Microbiol 57:244–245. 55. Neves FP, Ribeiro RL, Duarte RS, Teixeira LM, Merquior VL. 2009. Emergence of the vanA genotype among Enterococcus gallinarum isolates colonising the intestinal tract of patients in a university hospital in Rio de Janeiro, Brazil. Int J Antimicrob Agents 33:211–215. 56. Kawalec M, Kedzierska J, Gajda A, Sadowy E, Wegrzyn J, Naser S, Skotnicki AB, Gniadkowski M, Hryniewicz W. 2007. Hospital outbreak of vancomycin-resistant enterococci caused by a single clone of Enterococcus raffinosus and several clones of Enterococcus faecium. Clin Microbiol Infect 13:893–901. 57. Wilke WW, Marshall SA, Coffman SL, Pfaller MA, Edmund MB, Wenzel RP, Jones RN. 1997. Vancomycinresistant Enterococcus raffinosus: molecular epidemiology, species identification error, and frequency of occurrence in National resistance surveillance program. Diagn Microbiol Infect Dis 28:43–49. 58. National Nosocomial Infections Surveillance System. 2004. National Nosocomial Infections Surveillance (NNIS) System Report, data summary from January 1992 through June 2004, issued October 2004. Am J Infect Control 32:470–485. 59. Kramer A, Schwebke I, Kampf G. 2006. How long do nosocomial pathogens persist on inanimate surfaces? A systematic review. BMC Infect Dis 6:130–137. 60. Hospital Infection Control Practices Advisory Committee. 1995. Recommendations for preventing the spread of vancomycin resistance. Infect Control Hosp Epidemiol 16:105–113. 61. Robinson GL, Harris AD, Morgan DJ, Pineles L, Belton BM, Kristie Johnson J, The Benefits of Universal Gloving and Gowning (BUGG) Study Group. 2012. Survival of methicillin-resistant Staphylococcus aureus and vancomycin-resistant Enterococcus spp. for an extended period of transport. J Clin Microbiol 50:2466–2468. 62. Palladino S, Kay ID, Flexman JP, Boehm I, Costa AM, Lambert EJ, Christiansen KJ. 2003. Rapid detection of vanA and vanB genes directly from clinical specimens and enrichment broths by real-time multiplex PCR assay. J Clin Microbiol. 41:2483–2486. 63. Stamper PD, Cai M, Lema C, Eskey K, Carroll KC. 2007. Comparison of the BD GeneOhm VanR assay to culture for identification of vancomycin-resistant enterococci in rectal and stool specimens. J Clin Microbiol 45:3360–3365. 64. Young HL, Ballard SA, Roffey P, Grayson ML. 2007. Direct detection of vanB2 using the Roche LightCycler vanA/B detection assay to indicate vancomycin-resistant enterococcal carriage—sensitive but not specific. J Antimicrob Chemother 58:809–810. 65. Casalta JP, Gouriet F, Roux V, Thuny F, Habib G, Raoult D. 2009. Evaluation of the LightCycler® SeptiFast test in the rapid etiologic diagnostic of infectious endocarditis. Eur J Clin Microbiol Infect Dis 28:569–573. 66. Huckabee CM, Huskins WC, Murray PR. 2009. Predicting clearance of colonization with vancomycin-resistant enterococci and methicillin-resistant Staphylococcus aureus

418 n

67.

68.

69.

70.

71.

72.

73.

74.

75.

76.

77.

78.

BACTERIOLOGY

by use of weekly surveillance cultures. J Clin Microbiol 47:1229–1230. Bourdon N, Bérenger R, Lepoultier R, Mouet A, Lesteven C, Borgey F, Fines-Guyon M, Leclercq R, Cattior V. 2010. Rapid detection of vancomycin-resistant enterococci from rectal swabs by the Cepheid Xpert vanA/vanB assay. Diagn Microbiol Infect Dis 67:291–293. Usacheva EA, Ginocchio CC, Morgan M, Maglanoc G, Mehta MS, Tremblay S, Karchmer TB, Peterson LR. 2010. Prospective, multicenter evaluation of the BD GeneOhm VanR assay for direct, rapid detection of vancomycin-resistant Enterococcus species in perianal and rectal specimens. Am J Clin Pathol 134:219–226. Fang H, Ohlsson AK, Jian GX, Ullberg M. 2011. Screening for vancomycin-resistant enterococci: an efficient and economical laboratory-developed test. Eur J Clin Microbiol Infect Dis 31:261–265. Gazin M, Lammens C, Goossens H, Malhotra-Kumar S, MOSAR WP 2 S tu dy Te am . 2 01 1. Ev al uation of GeneOhm vanR and Xpert vanA/vanB molecular assays for the rapid detection of vancomycin-resistant enterococci. Eur J Clin Microbiol Infect Dis 31:273–276. Marner ES, Wolk DM, Carr J, Hewitt C, Dominguez LL, Kovacs T, Johnson DR, Hayden T. 2011. Diagnostic accuracy of the Cepheid GeneXpert assay ver.1.0 to detect the vanA and vanB Vancomycin resistance genes in Enterococcus from perianal specimens. Diagn Microbiol Infect Dis 69:382–389. Werner G, Serr A, Schutt S, Schneider C, Klare I, Witte W, Wendt C. 2011. Comparison of direct cultivation on a selective solid medium, polymerase chain reaction from an enrichment broth and the BD GeneOhm VanR assay for identification of vancomycin-resistant enterococci in screening specimens. Diagn Microbiol Infect Dis 70:512– 521. Zabicka D, Strzelecki J, Woznicak A, Strzelecki P, Sadowy E, Kuch A, Hryniewicz W. 2012. Efficiency of the Cepheid Xpert vanA/vanB assay for screening of colonization with vancomycin-resistant enterococci during hospital outbreak. Antonie van Leeuwenhoek 101:671–675. Babady NE, Gilhuley K, Clanciminio-Bordelon D, Tang YW. 2012. Performance characteristics of the Cepheid Xpert vanA assay for rapid identification of patients at high risk for carriage of vancomycin-resistant enterococci. J Clin Microbiol 50:3659–3663. Ballard SA, Grabsch FA, Johnson PDR, Grayson ML. 2006. Comparison of three PCR primer sets for identification of vanB gene carriage in feces and correlation with carriage of vancomycin-resistant enterococci: interference by vanB-containing anaerobic bacilli. Antimicrob Agents Chemother 49:77–81. Peters RP, van Agtmael MA, Gierveld S, Danner SA, Groeneveld AB, Vandenbroucke-Grauls CM, Savelkoul PH. 2007. Quantitative detection of Staphylococcus aureus and Enterococcus faecalis DNA in blood to diagnose bacteremia in patients in the intensive care unit. J Clin Microbiol 45:3641–3646. Weber SG, Huang SS, Oriola S, Huskins WC, Noskin GA, Harriman K, Olmsted RN, Bonten M, Lundstrom T, Climo MW, Roghmann MC, Murphy CL, Karchmer TB. 2007. Legislative mandates for use of active surveillance cultures to screen for methicillin-resistant Staphylococcus aureus and vancomycin-resistant enterococci: position statement from the joint SHEA and APIC task force. Infect Control Hosp Epidemiol 28:249–260. Delmas J, Robin F, Schweitzer C, Lesens O, Bonnet R. 2007. Evaluation of a new chromogenic medium chromID VRE, for detection of vancomycin-resistant enterococci in stool samples and rectal swabs. J Clin Microbiol 45:2731– 2735.

79. Park I, Park RW, Lim SK, Lee W, Shin JS, Yu S, Shin GT, Kim H. 2011. Rectal cultures screening for vancomycin-resistant Enterococcus in chronic haemodialysis patients; false-negative rates and duration of colonization. J Hosp Infect 79:147–150. 80. Kuch A, Strfaniuk E, Ozorowski T, Hryniewicz W. 2009. New selective and differential chromogenic agar medium, chromID VRE for screening vancomycin-resistant Enterococcus species. J Microbiol Methods 77:124–126. 81. Novicki TJ, Schapiro JM, Ulness BK, Sebeste A, BusseJohnston L, Swanson KM, Swanzy SR, Leisenring W, Limaye AP. 2004. Convenient selective differential broth for isolation of vancomycin-resistant Enterococcus from fecal material. J Clin Microbiol 42:1637–1640. 82. Brown FJ, Walpole E. 2003. Evaluation of selective enrichment media for isolation of glycopeptide-resistant enterococci from faecal specimens. J Antimicrob Chemother 51:289–296. 83. Ieven M, Vercauteren E, Descheemacker P, van Laer F, Goossens H. 1999. Comparison of direct plating and broth enrichment culture for detection of intestinal colonization by glycopeptide-resistant enterococci among hospitalized patients. J Clin Microbiol 37:1436–1440. 84. Seo JY, Kim PW, Lee JH, Song JH, Peck KR, Chung DR, Kang CI, Ki CS, Lee NY. 2011. Evaluation of PCRbased screening for vancomycin-resistant enterococci compared with a chromogenic agar-based culture method. J Med Microbiol 60:945–949. 85. Landman D, Quale JM, Oydna E, Wiley B, Ditore V, Zamam M, Patel K, Saurina G, Huang W. 1996. Comparison of five selective media for identifying fecal carriage of vancomycin-resistant enterococci. J Clin Microbiol 34:751– 753. 86. Paule SM, Trick WE, Tenover FC, Lankford M, Cunningham S, Stosor V, Cordell RL, Peterson LR. 2003. Comparison of PCR assay to culture for surveillance detection of vancomycin-resistant enterococci. J Clin Microbiol 41:4805–4807. 87. Chang JC, Chien ML, Chen HM, Yan JJ, Wu JJ. 2008. Comparison of CPS ID 3 and CHROMagar Orientation chromogenic agars with standard biplate technique for culture of clinical urine samples. J Microbiol Immunol Infect 41:422–427. 88. Graham M, Ballard SA, Grabsch EA, Johnson PDR, Grayson ML. 2008. High rates of fecal carriage of nonenterococcal vanB in both children and adults. Antimicrob Agents Chemother 52:1195–1197. 89. Klare I, Fleige C, Geringer U, Witte W, Werner G. 2012. Performance of three chromogenic VRE screening agars, two Etest vancomycin protocols, and different microdilution methods in detecting vanB genotype Enterococcus faecium with varying vancomycin MICs. Diagn Microbiol Infect Dis 74:171–176. 90. Ongut H, Kilinckaya H, Baysan GO, Ogunc D, Colak D, Inan D, Kasaroglu K, Gunseren F. 2013. Evaluation of Brillance VRE agar for the detection of vancomycinresistant enterococci in rectal swab speciments. J Med Microbiol 62:661–662. 91. Carvalho MGS, Steigerwalt AG, Morey RE, Shewmaker PL, Teixeira LM, Facklam RR. 2004. Characterization of three new enterococcal species, Enterococcus sp. nov. CDC PNS-E1, Enterococcus sp. nov. CDC PNS-E2, and Enterococcus sp. nov. CDC PNS-E3, isolated from human clinical specimens. J Clin Microbiol 42:1192–1198. 92. Carvalho MG, Steigerwalt AG, Morey RE, Shewmaker PL, Falsen E, Facklam RR, Teixeira LM. 2008. Designation of the provisional new Enterococcus species CDC PNSE2 as Enterococcus sanguinicola sp. nov., isolated from human blood, and identification of a strain previously named Enterococcus CDC PNS-E1 as Enterococcus italicus

23. Enterococcus n

93.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

Fortina, Ricci, Mora, and Manachini 2004. J Clin Microbiol 46:3473–3476. Garcia-Garrote F, Cercenado E, Bouza E. 2000. Evaluation of a new system, VITEK 2, for identification and antimicrobial susceptibility testing of enterococci. J Clin Microbiol 38:2108–2111. d’Azevedo PA, Dias CAG, Goncalves ALS, Rowe F, Teixeira LM. 2001. Evaluation of an automated system for the identification and antimicrobial susceptibility testing of enterococci. Diagn Microbiol Infect Dis 42:157–161. Brigante G, Luzzaro F, Bettaccini A, Lombarda G, Meacci F, Pini B, Stefani S, Toniolo A. 2006. Use of the Phoenix automated system for identification of Streptococcus and Enterococcus spp. J Clin Microbiol 44:3263– 3267. Clark AE, Kaleta EJ, Arora A, Wolk DM. 2013. Matrixassisted laser desorption ionization-time of flight mass spectrometry: a fundamental shift in the routine practice of clinical microbiology. Clin Microbiol Rev 26:547–603. van Belkum A, Welker M, Erhard M, Chatellier S. 2012. Biomedical mass spectrometry in today’s and tomorrow’s clinical microbiology laboratories. Clin Microbiol 50: 1513–1517. Fang H, Ohlsson AK, Ullberg M, Ozenci V. 2012. Evaluation of species-specific PCR, Bruker MS, VITEK MS and the VITEK 2 system for the identification of clinical Enterococcus isolates. Eur J Clin Microbiol Infect Dis 31:3073–3977. Quintela-Baluja M, Böhme K, Fernández-No IC, Morandi S, Alnakip ME, Caamaño-Antelo S, BarrosVelázquez J, Calo-Mata P. 2013. Characterization of different food-isolated Enterococcus strains by MALDI-TOF mass fingerprinting. Electrophoresis 34:2240–2250. Rychert J, Burnham CA, Bythrow M, Garner OB, Ginocchio CC, Jennemann R, Lewinski MA, Manji R, Mochon AB, Procop GW, Richter SS, Sercia L, Westblade LF, Ferraro MJ, Branda JA. 2013. Multicenter evaluation of the Vitek MS matrix-assisted laser desorption ionization-time of flight mass spectrometry system for identification of Gram-positive aerobic bacteria. J Clin Microbiol 51:2225–2231. Schulthess B, Brodner K, Bloemberg GV, Zbinden R, Böttger EC, Hombach M. 2013. Identification of Grampositive cocci by use of matrix-assisted laser desorption ionization-time of flight mass spectrometry: comparison of different preparation methods and implementation of a practical algorithm for routine diagnostics. J Clin Microbiol 51:1834–1840. McElvania Tekippe E, Shuey S, Winkler DW, Butler MA, Burnham CA. 2013. Optimizing identification of clinically relevant Gram-positive organisms by use of the Bruker Biotyper matrix-assisted laser desorption ionization-time of flight mass spectrometry system. J Clin Microbiol 51:1421–1427. March-Rosselló GA, Muñoz-Moreno MF, GarcíaLoygorri-Jordán de Urriés MC, Bratos-Pérez MA. 2013. A differential centrifugation protocol and validation criterion for enhancing mass spectrometry (MALDI-TOF) results in microbial identification using blood culture growth bottles. Eur J Clin Microbiol Infect Dis. 32:699– 704. Nori P, Ostrowsky B, Dorokhova O, Gialanella P, Moy M, Muggia V, Grossberg R, Kornblum J, Lin Y, Levi MH. 2013. Use of matrix-assisted laser desorption ionization-time of flight mass spectrometry to resolve complex clinical cases of patients with recurrent bacteremias. J Clin Microbiol 51:1983–1086. Domig KJ, Mayer HK, Kneifel W. 2003. Methods used for the isolation, enumeration, characterisation and identification of Enterococcus spp. 2. Pheno- and genotypic criteria. Int J Food Microbiol 88:165–188.

419

106. Merquior VLC, Peralta JM, Facklam RR, Teixeira LM. 1994. Analysis of electrophoretic whole-cell protein profiles as a tool for characterization of Enterococcus species. Curr Microbiol 28:149–153. 107. Teixeira LM, Facklam RR, Steigerwalt AG, Pigott NE, Merquior VLC, Brenner DJ. 1995. Correlation between phenotypic characteristics and DNA relatedness with Enterococcus faecium strains. J Clin Microbiol 33:1520– 1523. 108. Poyart C, Quesnes G, Trieu-Cuot P. 2000. Sequencing the gene encoding manganese-dependent superoxide dismutase for rapid species identification of enterococci. J Clin Microbiol 38:415–418. 109. Drancourt M, Roux V, Fournier PE, Raoult D. 2004. rpoB gene sequence-based identification of aerobic Grampositive cocci of the genera Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella. J Clin Microbiol 42:497–504. 110. Picard FJ, Ke D, Boudreau DK, Boissinot M, Huletsky A, Richard D, Ouellette M, Roy PH, Bergeron MG. 2004. Use of tuf sequences for genus-specific PCR detection and phylogenetic analysis of 28 streptococcal species. J Clin Microbiol 42:3686–3695. 111. Naser SM, Thompson FL, Hoste B, Gevers D, Dawyndt P, Vancanneyt M, Swings J. 2005. Application of multilocus sequence analysis (MLSA) for rapid identification of Enterococcus species based on rpoA and pheS genes. Microbiology 151:2141–2150. 112. Shewmaker PL, Steigerwalt AG, Nicholson AC, Carvalho Mda G, Facklam RR, Whitney AM, Teixeira LM. 2011. Reevaluation of the taxonomic status of recently described species of Enterococcus: evidence that E. thailandicus is a senior subjective synonym of “E. sanguinicola” and confirmation of E. caccae as a species distinct from E. silesiacus. J Clin Microbiol 49:2676–2679. 113. Sistek V, Maheux AF, Boissinot M, Bernard KA, Cantin P, Cleenwerck I, De Vos P, Bergeron MG. 2012. Enterococcus ureasiticus sp. nov. and Enterococcus quebecensis sp. nov., isolated from water. Int J Syst Evol Microbiol 62:1314–1320. 114. Depardieu F, Perichon B, Courvalin P. 2004. Detection of the van alphabet and identification of enterococci and staphylococci at the species level by multiplex PCR. J Clin Microbiol 42:5857–5860. 115. Forrest GN, Roghmann MC, Toombs LS, Johnson JK, Weekes E, Lincalis DP, Venezia RA. 2008. Peptide nucleic acid fluorescent in situ hybridization for hospitalacquired enterococcal bacteremia: delivering earlier effective antimicrobial therapy. Antimicrob Agents Chemother 52:3558–3563. 116. Gescher DM, Kovaecevic D, Schmiedel D, Siemoneit S, Mallmann C, Halle E, Gobel UB, Moter A. 2008. Fluorescence in situ hybridization (FISH) accelerates identification of Gram-positive cocci in positive blood cultures. Int J Antimicrob Agents 32S:S51–S59. 117. Morgan MA, Marlowe E, Novak-Weekly S, Miller JM, Painter TM, Salimnia H, Crystal B. 2011. A 1.5 hour procedure for identification of Enterococcus species directly from blood cultures. J Vis Exp. 48:pii:2616. doi:10.3791/2616. 118. Buchan BW, Ginocchio CC, Manii R, Cavagnolo R, Pancholi P, Swyers L, Thomson RB, Jr, Anderson C, Kaul K, Ledeboer NA. 2013. Multiplex identification of gram-positive bacteria and resistance determinants directly from positive blood culture broths: evaluation of an automated microarray-based nucleic acid test. PLoS Med 10:e1001478. 119. Scott LJ. 2013. Verigene® gram-positive blood culture nucleic acid test. Mol Diagn Ther 17:117–122. 120. Blaschke AJ, Heyrend C, Byington CL, Fisher MA, Barker E, Garrone NF, Thatcher SA, Pavia AT, Barney

420 n

121.

122.

123.

124.

125.

126.

127.

128.

129.

130.

131.

132.

133.

BACTERIOLOGY

T, Alger GD, Daly JA, Ririe KM, Ota I, Portiz MA. 2012. Rapid identification of pathogens from positive blood cultures by multiplex polymerase chain reaction using FilmArray system. Diagn Microbiol Infect Dis 74:349–355. Sango A, McCarter YS, Johnson D, Ferreira J, Guzman N, Jankowski CA. 2013. Stewardship approach for optimizing antimicrobial therapy through use of a rapid microarray assay on blood cultures positive for Enterococcus species. J Clin Microbiol 51:4008–4011. Freitas AR, Novais C, Ruiz-Garbajosa P, Coque TM, Peixe L. 2009.Clonal expansion within clonal complex 2 and spread of vancomycin-resistant plasmids among different genetic lineages of Enterococcus faecalis from Portugal. J Antimicrob Chemother 63:1104–1111. Valdezate S, Labayru C, Navarro A, Mantecón MA, Ortega M, Coque TM, García M, Saéz-Nieto JA. 2009. Large clonal outbreak of multidrug-resistant CC17 ST17 Enterococcus faecium containing Tn5382 in a Spanish hospital. J Antimicrob Chemother 63:17–20. Willems RJ, Top J, van Santen M, Robinson DA, Coque TM, Baquero F, Grundmann H, Bonten MJ. 2005. Global spread of vancomycin-resistant Enterococcus faecium from distinct nosocomial genetic complex. Emerg Infect Dis 11:821–828. Miranda AG, Singh KV, Murray BE. 1991. DNA fingerprinting of Enterococcus faecium by pulsed-field gel electrophoresis may be a useful epidemiologic tool. J Clin Microbiol 29:2752–2757. Kawalec M, Gniadkowski M, Hryniewicz W. 2000. Outbreak of vancomycin-resistant enterococci in a hospital in Gdansk, Poland, due to horizontal transfer of different Tn1546-like transposon variants and clonal spread of several strains. J Clin Microbiol 38:3317–3322. Tenover FC, Arbeit RD, Goering RV, Mickelsen PA, Murray BE, Persing DH, Swaminathan B. 1995. Interpreting chromosomal DNA restriction patterns produced by pulsed-field gel electrophoresis: criteria for bacterial strain typing. J Clin Microbiol 33:2233–2239. Homan WL, Tribe D, Poznanski S, Li M, Hogg G, Spalburg E, Van Embden JD, Willems RJ. 2002. Multilocus sequence typing scheme for Enterococcus faecium. J Clin Microbiol 40:1963–1971. Ruiz-Garbajosa P, Bonten MJM, Robinson DA, Top J, Nallapareddy SR, Torres C, Coque TM, Canton R, Baquero F, Murray BE, del Campo R, Willems RJ. 2006. A multilocus sequence typing scheme for Enterococcus faecalis reveals hospital-adapted genetic complexes in a background of high rates of recombination. J Clin Microbiol 44:2220–2228. Werner G, Coque TM, Hammerum AM, Hope R, Hryniewicz W, Johnson A, Klare I, Kristinsson KG, Leclercq R, Lester CH, Lillie M, Novais C, OlssonLiljequist B, Peixe LV, Sadowy E, Simonsen GS, Top J, Vuopio-Varkila J, Willems RJ, Witte W, Woodford N. 2008. Emergence and spread of vancomycin resistance among enterococci in Europe. Euro Surveill 13:1–11. Galloway-Peña J, Roh JH, Latorre M, Qin X, Murray BE. 2012. Genomic and SNP analyses demonstrate a distant separation of the hospital and community-associated clades of Enterococcus faecium. PLoS One 7:e30187. Titze-de-Almeida R, Willems RJ, Top J, Rodrigues IP, Ferreira RF, II, Boelens H, Brandileone MC, Zanella RC, Felipe MSS, van Belkum A. 2004. Multilocus variable-number tandem-repeat polymorphism among Brazilian Enterococcus faecalis strains. J Clin Microbiol 42:4879–4881. Top J, Schouls LM, Bonten MJ, Willems RJ. 2004. Multiple-locus variable-number tandem repeat analysis, a novel typing scheme to study the genetic relatedness

134.

135.

136.

137.

138.

139.

140.

141. 142.

143.

144.

145.

146.

147.

148. 149.

150.

and epidemiology of Enterococcus faecium isolates. J Clin Microbiol 42:4503–4511. Top J, Banga NM, Hayes R, Willems RJ, Bonten MJ, Hayden MK. 2008. Comparison of multiple-locus variable-number tandem repeat analysis and pulsed-field gel electrophoresis in a setting of polyclonal endemicity of vancomycin-resistant Enterococcus faecium. Clin Microbiol Infect 14:363–369. Didelot X, Bowden R, Wilson DJ, Peto TE, Crook DW. 2012. Transforming clinical microbiology with bacterial genome sequencing. Nat Rev Genet 13:601–612. Reuter S, Ellington MJ, Cartwright EJ, Köser CU, Török ME, Gouliouris T, Harris SR, Brown NM, Holden MT, Quail M, Parkhill J, Smith GP, Bentley SD, Peacock SJ. 2013. Rapid bacterial whole-genome sequencing to enhance diagnostic and public health microbiology. JAMA Intern Med 173:1397–1404. Zankari E, Hasman H, Kaas RS, Seyfarth AM, Agersø Y, Lund O, Larsen MV, Aarestrup FM. 2013. Genotyping using whole-genome sequencing is a realistic alternative to surveillance based on phenotypic antimicrobial susceptibility testing. J Antimicrob Chemother 68:771–777. Cho YS, Lee HS, Kim JM, Lee MH, Yoo HS, Park YH, Ryu PD. 2008. Immunogenic proteins in the cell envelope and cytoplasm of vancomycin-resistant enterococci. J Immunoassay Immunochem 29:319–331. Sulaiman A, Rakita RM, Arduino RC, Patterson JE, Steckelberg JM, Singh KV, Murray BE. 1996. Serological investigation of enterococcal infections using western blot. Eur J Clin Microbiol Infect Dis 15:826–829. Kak V, Chow JW. 2002. Acquired antibiotic resistances in enterococci, p 355–383. In Gilmore MS, Clewell DB, Courvalin P, Dunny GM, Murray BE, Rice LB (ed), The Enterococci: Pathogenesis, Molecular Biology, and Antibiotic Resistance. ASM Press, Washington, DC. Courvalin P. 2006. Vancomycin resistance in grampositive cocci. Clin Infect Dis 42:S25–S34. Hollenbeck BL, Rice LB. 2012. Intrinsic and acquired resistance mechanisms in Enterococcus. Virulence 3:421– 433. Linden PK. 2007. Optimizing therapy for vancomycinresistant enterococci (VRE). Semin Respir Crit Care Med 28:632–645. Arias CA, Contreras GA, Murray BE. 2010. Management of multidrug-resistant enterococcal infections. Clin Microbiol Infect 16:555–562. Clinical and Laboratory Standards Institute. 2014. Performance Standards for Antimicrobial Susceptibility Testing. CLSI Document M100-S24, Twenty-Fourth Informational Supplement. Clinical and Laboratory Standards Institute, Wayne, PA. Weinstein MP. 2001. Comparative evaluation of penicillin, ampicillin, and imipenem MICs and susceptibility breakpoints for vancomycin-susceptible and vancomycinresistant Enterococcus faecalis and Enterococcus faecium. J Clin Microbiol 39:2729–2731. Leclercq R, Derlot E, Duval J, Courvalin P. 1988. Plasmid-mediated resistance to vancomycin and teicoplanin in Enterococcus faecium. N Engl J Med 319:157– 161. Uttley AHC, Collins CH, Naidoo J, George RC. 1988. Vancomycin-resistant enterococci. Lancet i:57–58. Clark NC, Teixeira LM, Facklam RR, Tenover FC. 1998. Detection and differentiation of the vanC-1, vanC2, and vanC-3 glycopeptide resistance genes in enterococci. J Clin Microbiol 36:2294–2297. Watanabe S, Kobayashi N, Quiñones D, Hayakawa S, Nagashima S, Uehara N, Watanabe N. 2009. Genetic diversity of the low-level vancomycin resistance gene vanC-2/vanC-3 and identification of a novel vanC subtype

23. Enterococcus n (vanC-4) in Enterococcus casseliflavus. Microb Drug Resist 15:1–9. 151. Boyd DA, Willey BM, Fawcett D, Gillani N, Mulvey MR. 2008. Molecular characterization of Enterococcus faecalis N06-0364 with low-level vancomycin resistance harboring a novel D-Ala-D-Ser gene cluster, vanL. Antimicrob Agents Chemother 52:2667–2672. 152. Xu X, Lin D, Yan G, Ye X, Wu S, Guo Y, Zhu D, Hu F, Zhang Y, Wang F, Jacoby GA, Wang M. 2010. vanM, a new glycopeptide resistance gene cluster found in Enterococcus faecium. Antimicrob Agents Chemother 54:4643– 4647. 153. Lebreton F, Depardieu F, Bourdon N, Fines-Guyon M, Berger P, Camiade S, Leclercq R, Courvalin P, Cattoir

421

V. 2011. D-Ala-d-Ser VanN-type transferable vancomycin resistance in Enterococcus faecium. Antimicrob Agents Chemother 55:4606–4612. 154. Tambyah PA, Marx JA, Maki DG. 2004. Nosocomial infection with vancomycin-dependent enterococci. Emerg Infect Dis 10:1277–1281. 155. Alam MR, Donabedian S, Brown W, Gordon J, Chow JW, Zervos MJ, Hershberger E. 2001. Heteroresistance to vancomycin in Enterococcus faecium. J Clin Microbiol 39:3379–3381. 156. Garnica ML, Valdezate S, Gonzalo C, Saez-Nieto JA. 2013. Presence of the vanC1 gene in a vancomycin-resistant Enterococcus faecalis strain isolated from ewe bulk tank milk. J Med Microbiol 62:494–495.

Aerococcus, Abiotrophia, and Other Aerobic Catalase-Negative, Gram-Positive Cocci JENS JØRGEN CHRISTENSEN AND KATHRYN L. RUOFF

24 Abiotrophia elegans, was described in 1998 (9). Kanamoto et al. noted the heterogeneity among Abiotrophia strains and proposed a fourth species, Abiotrophia para-adiacens (10). In 2000, Collins and Lawson proposed a new genus, Granulicatella, with Granulicatella adiacens and Granulicatella elegans representing strains formerly called A. adiacens and A. elegans. A. defectiva remains as the sole Abiotrophia species (6). Among the intrinsically vancomycin-resistant, catalasenegative, Gram-positive cocci, a number of Leuconostoc species have been noted in human infection (Leuconostoc mesenteroides, Leuconostoc lactis, Leuconostoc pseudomesenteroides, and Leuconostoc citreum [11]). In 1993, the former Leuconostoc paramesenteroides and related species were placed into a novel genus, Weissella (12). Pediococcus acidilactici and Pediococcus pentosaceus are the most common clinical isolates of pediococci (13). The vancomycin-susceptible species formerly named Pediococcus halophilus was reclassified in the genus Tetragenococcus (14). The organism formerly called Enterococcus solitarius has also been transferred to the Tetragenococcus genus as Tetragenococcus solitarius (15). Little is known about the role of the tetragenococci in human infection. The organism we now know as Gemella morbillorum was described in 1917 by Tunnicliff (16) as an isolate from the blood of patients with measles. G. morbillorum was originally named Diplococcus rubeolae and was also called Diplococcus morbillorum, Peptostreptococcus morbillorum, and Streptococcus morbillorum until a proposal to include it in the genus Gemella as G. morbillorum was made in 1988 (17). A second species, Gemella haemolysans, was originally classified as a Neisseria species, due to its Gram-variable or even Gramnegative nature and its cellular morphology (diplococci with flattened adjacent sides). Collins and coworkers described two additional Gemella species isolated from human sources, Gemella bergeri (originally named Gemella bergeriae [18]) and Gemella sanguinis (19). Gemella asaccharolytica was recently described by Ulger-Toprak and colleagues (20). The genus Dolosigranulum shows phenotypic similarities to Gemella, although it is not phylogenetically closely related to Gemella strains (21, 22). Aerococcus urinae, described in 1992, is negative for pyrrolidonyl arylamidase production (PYR) and positive for leucine aminopeptidase production (LAP), showing opposite reactions of Aerococcus viridans in these important identification tests (23). In spite of these phenotypic differences,

TAXONOMY The catalase-negative, Gram-positive cocci included in this chapter form a taxonomically diverse group of bacteria that are isolated infrequently as opportunistic agents of infection. Most of these organisms resemble other more well-known clinical isolates (i.e., streptococci and enterococci) and consequently may be mistaken for members of those genera. Although probably misidentified or overlooked in clinical cultures in the past, these organisms may represent emerging pathogens in immunocompromised patient populations. Table 1 lists the organisms included here along with some of their basic characteristics. The bacteria discussed in this chapter are members of the phylum Firmicutes (low-G+C, Gram-positive bacteria). Helcococcus is the only genus in the group to reside in the class “Clostridia,” while the remaining genera are classified in the class “Bacilli” (1). The reader is referred to chapter 21 for information on Rothia mucilaginosa, another infrequently isolated Gram-positive coccus that may be catalase negative. The genus Lactococcus is composed of organisms formerly classified as Lancefield group N streptococci (2). The species Lactococcus lactis and Lactococcus garvieae have been documented in human infections. Motile Lactococcus-like organisms with Lancefield’s group N antigen (a teichoic acid antigen) are classified in the genus Vagococcus (3, 4). The vagococci also resemble the enterococci, and Facklam and Elliott (5) reported that Vagococcus fluvialis (the principal species described in human clinical specimens to date) isolates examined at the Centers for Disease Control and Prevention gave positive reactions in a commercially available nucleic acid probe test for enterococci. The genera Abiotrophia and Granulicatella accommodate organisms previously known as nutritionally variant or satelliting streptococci (6, 7). These bacteria were originally thought to be nutritional mutants of viridans streptococcal strains, most notably of the species Streptococcus mitis. Bouvet and colleagues (8) suggested that this group of organisms were really members of two novel streptococcal species given the names Streptococcus defectivus and Streptococcus adjacens. A comparative analysis of 16S rRNA gene sequences led Kawamura and coworkers to propose the creation of a new genus, Abiotrophia, containing two species, Abiotrophia defectiva and Abiotrophia adiacens, to accommodate these bacteria (7). A third species from human sources,

doi:10.1128/9781555817381.ch24

422

24. Aerobic Catalase-Negative, Gram-Positive Cocci n

423

TABLE 1 Possible identities of catalase-negative, Gram-positive cocci based on certain phenotypic reactions and cellular morphologya Phenotypic reaction PYR

LAP

NaCl

+

+

+

+

+

−

+ + − −

− − + +

+ − + −

−

+

−

Cellular morphology Pairs, chains

Clusters, tetrads, irregular groups

Enterococcus,b,c,d Vagococcus,c,e Lactococcus,d Facklamia spp. other than F. languida, Ignavigranumf Abiotrophia,g Granulicatella,g Gemella spp. other than G. haemolysans and G. asaccharolytica Globicatella Dolosicoccus

F. languida, Dolosigranulum, Aerococcus sanguinicola G. haemolysans (cells arranged primarily in pairs, with adjacent sides flattened) Aerococcus viridans,h Helcococcus kunziih,i Aerococcus urinae, Pediococcus,j Tetragenococcus

Viridans group streptococci,k G. asaccharolytica (Gram variable) Leuconostoc,j Weissellaj

a

Abbreviations and symbols: NaCl, growth in 6.5% NaCl; +, ≥90% of strains positive; −, ≤10% of strains positive. Some strains may display vancomycin resistance, and some strains are motile. Most enterococcal strains are capable of growth at 45°C, differentiating them from vagococci, which may be phenotypically similar. Strains of vagococci have been reported as testing positive with a commercially available nucleic acid probe for members of the genus Enterococcus. d Phenotypically similar strains of enterococci and lactococci can be differentiated with a commercially available nucleic acid probe for members of the genus Enterococcus. e Motile. f Some strains display satelliting growth; some strains are urease positive. g Members of this genus display satelliting growth. h Although H. kunzii shares some phenotypic traits with A. viridans, it is facultative and usually nonhemolytic, in contrast to A. viridans, which prefers an aerobic growth atmosphere and is alpha-hemolytic. i Two additional species of Helcococcus (H. sueciensis and “H. pyogenes”) have been proposed (36–38), both based on the isolation of a single strain. These new species display negative reactions in the PYR test, in contrast to H. kunzii. j Vancomycin resistant. k Viridans group streptococci include streptococci of the anginosus, mitis, mutans, salivarius, and bovis species groups. Some strains of Streptococcus pneumoniae (a member of the mitis species group) may produce positive reactions in the PYR test. b c

molecular taxonomic studies suggest that A. urinae should remain in the Aerococcus genus. Organisms currently included in the A. urinae species are fairly heterogeneous and can probably be subdivided into at least two subspecies (24). Aerococcus christensenii, isolated from the human genitourinary tract, was described by Collins and coworkers in 1999 (25) and was joined by the species Aerococcus sanguinicola (originally named Aerococcus sanguicola [26, 27]) and Aerococcus urinaehominis (28) in 2001. Globicatella, Facklamia, Ignavigranum, and Dolosicoccus are related genera that are isolated infrequently from clinical specimens. Globicatella sanguinis, initially named Globicatella sanguis, was described in 1992 (29). Facklamia currently contains four species isolated from human sources: Facklamia hominis (30), Facklamia sourekii (31), Facklamia ignava (32), and Facklamia languida (33). The genus Ignavigranum, currently consisting of a single species, Ignavigranum ruoffiae, was described by Collins and coworkers (34), along with the genus Dolosicoccus and its single species, Dolosicoccus paucivorans (35). The genus Helcococcus, originally composed of the single species Helcococcus kunzii (36), came to include a new species isolated from humans, Helcococcus sueciensis, in 2004 (37). A third human species, “Helcococcus pyogenes,” has been proposed, but to date has not received official taxonomic standing (38, 39). Helcococcus ovis, isolated from infections in animals, displays satelliting growth, unlike the human Helcococcus species (40).

DESCRIPTION OF THE GENERA The organisms included in this chapter form Gram-positive coccoid cells, but G. haemolysans may appear Gram variable or Gram negative due to the ease with which its cells are decolorized. Cell shape and arrangement can be used to

divide these organisms into two broad groups: those with a “streptococcal-like” Gram stain (coccobacilli in pairs and chains) and those with a “staphylococcal-like” Gram stain (more spherical cocci in pairs, tetrads, clusters, or irregular groups). Abiotrophia and Granulicatella isolates (formerly the nutritionally variant streptococci) form coccobacilli arranged in pairs and chains, but these organisms may also appear pleomorphic, especially when grown under suboptimal nutritional conditions (41). Dividing these diverse bacteria into two groups based on cellular shape and arrangement serves only as an aid in identification; no relatedness of organisms is implied by this grouping. With the exception of the infrequently isolated vagococci, these bacteria are all nonmotile. Most of the genera described here are catalase-negative facultative anaerobes, but A. viridans is classified as a microaerophile that grows poorly, if at all, under anaerobic conditions. Some strains of Aerococcus may exhibit weakly positive catalase reactions due to nonheme catalase activity. None of the genera are beta-hemolytic on routinely employed blood agars, but strains of G. haemolysans, G. bergeri, and Gemella sanguinis have been described as beta-hemolytic on agars supplemented with horse blood (18, 19, 42).

EPIDEMIOLOGY AND TRANSMISSION The organisms discussed in this chapter are opportunistic pathogens. Some of the genera have been characterized as constituents of the normal microbiota of the human oral cavity or upper respiratory tract (Gemella, Abiotrophia, and Granulicatella) and skin (Helcococcus). Lactococci, pediococci, and leuconostocs can be isolated from foods and vegetation (5, 43, 44) and may also be found as part of the normal microbiota of the alimentary tract. Aerococci are environmental isolates that can also be found on human

424 n

BACTERIOLOGY

skin. Although they have been isolated from human clinical cultures, the natural habitats of many of the organisms mentioned here are not well characterized. The bacteria examined here seem to be of low virulence and are usually pathogenic only in immunocompromised hosts. Infection often occurs in previously damaged tissues (e.g., heart valves) or may be nosocomial and associated with prolonged hospitalization, antibiotic treatment, invasive procedures, and the presence of foreign bodies.

CLINICAL SIGNIFICANCE The bacteria described in this chapter may be present as contaminants in clinical cultures, but they are also isolated infrequently as opportunistic pathogens. Blood, cerebrospinal fluid, urine, and wound specimens are likely to yield significant isolates of these bacteria. Details on reported infections due to each of the genera follow.

Lactococcus Due to their phenotypic similarities with streptococci and enterococci, clinical isolates of lactococci have probably been misidentified in the past, accounting at least in part for the paucity of reports concerning the clinical role of these bacteria. Elliott and coworkers (45) studied the phenotypic characteristics of a number of lactococcal strains isolated from blood, urinary tract infections, and an eye wound culture. Lactococci have been associated with native valve and prosthetic valve endocarditis (46–49), septicemia in an immunosuppressed patient, osteomyelitis, spondylodiscitis (46), peritonitis (50), liver abscess (51, 52), acalculous cholecystitis (53), and prosthetic joint infection (54). L. garvieae is a known pathogen of aquacultured fish, and human infections have been linked to consumption of or contact with fish (46, 54).

Vagococcus To date, only a handful of Vagococcus isolates from human sources have been reported in the literature. Teixeira and coworkers (55) described strains isolated from blood, peritoneal fluid, and a wound. Al-Ahmad and colleagues reported isolation of V. fluvialis from an infected root canal system (56). Vagococci are motile organisms that, like lactococci, elaborate Lancefield’s group N antigen (5). Difficulties encountered in identifying vagococci may partially account for their infrequent recognition in clinical cultures.

Abiotrophia and Granulicatella Organisms in the genera Abiotrophia and Granulicatella (formerly known as nutritionally variant streptococci) are normal residents of the oral cavity and are recognized as agents of endocarditis involving both native and prosthetic valves (57–60). These organisms have also been isolated from other types of infections, including ophthalmic infections (61, 62), central nervous system infections (63, 64), peritonitis in patients undergoing continuous ambulatory peritoneal dialysis (65), musculoskeletal infection (66), septic arthritis (67), and a breast implant-associated infection (68).

Leuconostoc, Pediococcus, and Weissella The vancomycin-resistant genera Leuconostoc and Pediococcus were first recognized in clinical specimens in the mid1980s. Handwerger and colleagues (69) observed that host defense impairment, invasive procedures breaching the integument, gastrointestinal symptoms, and prior antibiotic treatment were common features among adult patients with Leuconostoc infection. They also noted a predisposition to

Leuconostoc bacteremia among neonates, suggesting that infants may become colonized during delivery by leuconostocs inhabiting the maternal genital tract. Leuconostocs have been isolated from blood, cerebrospinal fluid, peritoneal dialysate fluid, and wounds. Case reports have implicated leuconostocs as agents of infection in osteomyelitis (70), ventriculitis (71), brain abscess (72), postsurgical endophthalmitis (73), and bacteremia in the setting of short gut syndrome. Short gut syndrome favors a microbiota with a high prevalence of Lactobacillus and Leuconostoc bacteria (74). Several reports have related Leuconostoc bacteremic episodes to the presence of short gut syndrome, central venous catheters, and disrupted bowel mucosa (75). Pediococcus strains have been isolated from bacteremia and cases of sepsis and hepatic abscess in compromised patients (13, 76–79). Barros and coworkers (13) noted that P. acidilactici was isolated from clinical specimens more frequently than P. pentosaceus and was also more commonly isolated from cases of bacteremia. Barton and coworkers noted the role of Pediococcus in bacteremia in infants with gastrointestinal malformations requiring surgical correction (76). Weissella confusa, formerly classified as Lactobacillus confusus, has been reported infrequently as an agent of bacteremia and endocarditis (80, 81).

Gemella G. haemolysans has been isolated from cases of endocarditis (82), meningitis (83), brain abscess (84), a total knee arthroplasty (85), and ocular infection (86–88). G. morbillorum has been implicated in cases of endocarditis (89, 90), empyema and lung abscess (91), septic shock (92), brain abscess (93), osteomyelitis (94), septic arthritis (95), and peritonitis (96). Information on the clinical significance of the other Gemella species continues to accumulate. G. bergeri and G. sanguinis have been isolated from blood cultures, and they may also be causative agents of endocarditis (18, 19, 97). A strain of G. sanguinis was isolated from an infected prosthetic hip joint (98), and G. asaccharolytica has been isolated from wound cultures (20).

Dolosigranulum Dolosigranulum, a genus phenotypically similar, but not closely related, to Gemella (21), has been documented in blood, eye, and respiratory specimens (22). The single species of the genus, Dolosigranulum pigrum, has been associated with nosocomial pneumonia and septicemia (99), synovitis (100), acute cholecystitis accompanied by acute pancreatitis (101), and biomaterial-associated arthritis (102).

Aerococcus A. viridans has been noted as a contaminant in clinical cultures and infrequently as a clinically significant isolate from cases of endocarditis and bacteremia and a case of spondylodiscitis (103–105). Four additional Aerococcus species isolated from humans have been described since the early 1990s. A. urinae (23, 106) has been implicated as a urinary tract pathogen in patients predisposed to infection (107–109) and as an agent of endocarditis (110, 111), lymphadenitis (112), and peritonitis (113). A. sanguinicola has been isolated from blood and urine specimens (26, 27) and cases of urosepsis and endocarditis (114). Little is currently known about the clinical significance of A. christensenii (isolated from vaginal specimens [25]) and A. urinaehominis (isolated from urine [28]).

24. Aerobic Catalase-Negative, Gram-Positive Cocci n 425

Globicatella

ISOLATION PROCEDURES

G. sanguinis, isolated from human clinical specimens, has been implicated in cases of bacteremia, urinary tract infection, and meningitis (29, 115, 116). A second species in the genus, Globicatella sulfidifaciens, has been isolated from purulent infections in domestic mammals (117).

Generally, there are no special requirements for isolation of the group of bacteria discussed here; general recommendations for the culture of blood, body fluids, and other specimens should be followed (see chapter 18). These organisms are likely to be isolated on rich, nonselective media (e.g., blood or chocolate agar and thioglycolate broth) since they are nutritionally fastidious. If selective isolation of the vancomycin-resistant genera Leuconostoc and Pediococcus is desired, Thayer-Martin medium may be used to inhibit normal microbiota or other contaminating microorganisms (127). Some of the genera (e.g., Helcococcus) grow slowly, forming tiny colonies that may not be visible unless extended incubation (48 to 72 h) is employed. The recovery of many of the genera included in this chapter may be enhanced by CO2 enrichment of the incubation atmosphere. Members of the genera Abiotrophia and Granulicatella usually grow on chocolate agar, on brucella agar with 5% horse blood, and in thioglycolate broth, but not on Trypticase soy agar with 5% sheep blood. These organisms can be cultured on nonsupportive media that have been appropriately supplemented (see “Procedures for Phenotypic Differentiation, Abiotrophia and Granulicatella” below).

Facklamia The Facklamia genus is closely related to, but phenotypically and phylogenetically distinct from, Globicatella (30). Strains of the four Facklamia species isolated from humans have been recovered from blood, wound, and genitourinary sites (30–33) and a case of chorioamnionitis (118).

Ignavigranum A limited number of isolates of I. ruoffiae, the sole species of Ignavigranum, have been described to date. Sites of isolation include a wound and an ear abscess (34).

Dolosicoccus The single species of the genus Dolosicoccus, D. paucivorans, has been isolated from blood cultures (35, 119).

Helcococcus H. kunzii can be isolated from intact skin of the lower extremities (120) as well as from mixed cultures of wounds, notably foot infections (36, 121). In such scenarios the clinical significance of this organism is difficult to interpret, since it may be present merely as a colonizer of the wound site. The ability of H. kunzii to function as an opportunist is, however, suggested by its isolation as the sole or predominant organism from an infected sebaceous cyst (122), a breast abscess (123), a postsurgical foot abscess (124), an infected prosthetic joint (125), and cases of bacteremia and empyema in intravenous drug users (126). Two additional species isolated from humans, H. sueciensis and “H. pyogenes,” are based on single isolates from a wound and a prosthetic joint infection, respectively (37–39).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS No special requirements for collection and transport of specimens for isolation of the organisms discussed in this chapter have been described. Routine procedures for collection, transport, and storage of specimens for aerobic culture allow for the isolation of these bacteria, since the majority are facultative anaerobes or microaerophiles. These organisms should also be recovered from specimens that have been collected and transported under anaerobic conditions (see chapter 18).

IDENTIFICATION Procedures for Phenotypic Differentiation While molecular characterization may be required for accurate species-level identification of the aerobic catalasenegative, Gram-positive cocci encountered infrequently in clinical laboratories, phenotypic methods can be helpful in characterization of these bacteria to the genus level. Gram stain morphology has been employed as a major decision point in the identification protocols in Fig. 1 and 2 and Table 1, with two general categories: morphology resembling that of streptococci, meaning cocci or coccobacilli in pairs and chains, versus staphylococcal morphology, consisting of coccoid cells arranged in pairs, clusters, tetrads, or irregular groups. Broth-grown cells (thioglycolate broth is suitable) should be used for making accurate morphological determinations. Note that Gemella and Facklamia strains may display either type of cellular morphology, depending on the species. Figures 1 and 2 display phenotypic tests used to differentiate the genera of bacteria discussed in this chapter. Descriptions of tests for catalase, PYR, LAP, βglucuronidase, and hippurate hydrolysis, as well as bile esculin agar and Lactobacillus MRS (DeMan, Rogosa, Sharpe) broth media, can be found in chapter 19 and reference 5. Additional phenotypic tests are described below in the discussion of identification criteria for each genus.

Lactococcus and Vagococcus

DIRECT EXAMINATION The organisms described in this chapter can be visualized in direct Gram stains of clinical material but have no outstanding morphological characteristics that distinguish them from commonly isolated Gram-positive cocci (streptococci and staphylococci). Although Abiotrophia and Granulicatella isolates may appear pleomorphic in direct Gram stains, they form Gram-positive cocci in pairs and chains when grown on nutritionally adequate media. Direct detection of these genera by antigenic methods has not been described, but some authors have employed amplification of 16S rRNA genes for direct detection in clinical specimens (59).

The members of the genera Lactococcus and Vagococcus are usually PYR and LAP positive, grow in the presence of 6.5% NaCl, and can be confused with enterococci or streptococci. For the salt tolerance test, heart infusion broth supplemented with 6.0% NaCl (producing a final NaCl concentration of 6.5%), with or without the acid-base indicator bromcresol purple, is inoculated with two or three colonies and incubated at 35°C for up to 72 h. Turbidity with or without a color change from purple to yellow indicates growth (5, 128). Facklam and colleagues (5, 128) recommended growth temperature tests for distinguishing lactococci from streptococci and enterococci. Consult Fig. 1 for growth temperature characteristics of each of the genera. For growth temperature tests, broths (heart infusion broth containing 1% glucose

426 n

BACTERIOLOGY

FIGURE 1 Identification of catalase-negative, Gram-positive cocci that grow aerobically with cells arranged in pairs and chains. Abbreviations: 6.5% NaCl, growth in broth containing 6.5% NaCl; bile esculin, hydrolysis of esculin in the presence of 40% bile; motility, motility in motility test medium; 45°C, growth at 45°C; 10°C, growth at 10°C; probe, reaction with commercially available nucleic acid probe for the genus Enterococcus; HIP, hydrolysis of hippurate; satellitism, satelliting growth behavior; ARG, arginine hydrolysis activity; BGUR, β-glucuronidase activity. doi:10.1128/9781555817381.ch24.f1

and bromcresol purple indicator) are inoculated with a single colony or drop of broth culture of the test strain and incubated at 35°C for up to 7 days. A water bath is recommended for incubation of cultures at 45°C. Turbidity with or without a change in the broth’s indicator to yellow indicates a positive test. The motile vagococci can be distinguished from lactococci with modified motility test medium, stab-inoculated and incubated at 30°C for up to 48 h, according to the method of Facklam and Elliott (5). Further information on the phenotypic traits of Lactococcus and Vagococcus isolates may be found in references 2, 45, 55, and 129.

Abiotrophia and Granulicatella A test for satelliting behavior is important for identification of these two genera. The strain to be examined is streaked for confluent growth on a medium that does not support growth or supports only weak growth (e.g., sheep blood agar). A single cross streak of Staphylococcus aureus (ATCC 25923 or another suitable strain) is applied to the inoculated

area. After incubation at 35°C in an atmosphere containing elevated CO2, strains of Abiotrophia or Granulicatella grow only in the vicinity of the staphylococcal growth. Some strains of Ignavigranum may also show satelliting behavior (34). Alternatively, media can be supplemented with pyridoxal. An aqueous stock solution of filter-sterilized 0.01% pyridoxal hydrochloride (which can be stored frozen) should be added to media to achieve a final concentration of 0.001%. Pyridoxal disks (Remel, Lenexa, KS) may also be used in the satelliting test. Detailed phenotypic information for the PYR- and LAPpositive Abiotrophia and Granulicatella species can be found in references 6, 41, 130, and 131. Davis and Peel (132) reported that the API 20 Strep system (bioMérieux, Durham, NC) was superior to the Rapid ID32 Strep system (bioMérieux) for identification of these organisms.

Leuconostoc, Pediococcus, and Weissella Members of the PYR-negative, vancomycin-resistant genera Leuconostoc, Pediococcus, and Weissella produce small, alpha-

24. Aerobic Catalase-Negative, Gram-Positive Cocci n

427

FIGURE 2 Identification of catalase-negative, Gram-positive cocci that grow aerobically with cells arranged in pairs, tetrads, clusters, or irregular groups. Abbreviations: 6.5% NaCl, growth in broth containing 6.5% NaCl; esculin, hydrolysis of esculin; BGUR, β-glucuronidase activity. doi:10.1128/9781555817381.ch24.f2

hemolytic or nonhemolytic colonies on blood agar. Vancomycin resistance can be tested by streaking several colonies over half of a Trypticase soy agar with 5% sheep blood plate. After placing a 30-μg vancomycin disk in the center of the inoculated area, the plate is incubated overnight in a CO2-enriched atmosphere at 35°C. Any zone of inhibition indicates susceptibility, while resistant strains exhibit no inhibition zone (5, 128). In addition to differing cellular morphologies (Table 1), these vancomycin-resistant genera, along with vancomycin-resistant strains of lactobacilli that form short coccoid cells, can be differentiated by tests for gas production from glucose and arginine hydrolysis. Leuconostocs produce gas and are always arginine negative. Lactobacilli are variable in both tests, but a positive arginine test for a gas-producing strain would rule out identity of the organism as a leuconostoc. Pediococci are gas production negative and show variable reactions in the arginine test, although P. acidilactici and P. pentosaceus, the two species commonly found in clinical material, are arginine positive. Weissella strains may be misidentified as leuconostocs or lactobacilli. These organisms produce gas from glucose. The few clinical isolates reported in the literature have been described as positive for hydrolysis of arginine (12, 81). MRS broth (BD Diagnostic Systems, Franklin Lakes, NJ; Hardy Diagnostics, Santa Maria, CA; see chapter 19), sealed with melted petrolatum and incubated for up to 7 days at 35°C, is used to test for gas production, indicated by displacement of the petrolatum plug (5, 128). The arginine hydrolysis test can be performed with Moeller decarboxylase broth containing arginine (5). Lancefield group D antigen

can be detected in pediococci (128). References 5, 11, 13, 128, and 131 should be consulted for further information on identification of Leuconostoc and Pediococcus to the species level.

Gemella On sheep blood agar media, members of the Gemella genus (usually PYR positive) form small colonies that are similar in appearance to those of viridans group streptococci. Slow growth of some Gemella strains may lead to confusion of these organisms with Abiotrophia or Granulicatella (formerly called nutritionally variant streptococci). A test for satelliting behavior should separate these two groups of bacteria (128), but Leung and coworkers (98) reported on a single G. sanguinis strain that exhibited “pseudosatelliting” behavior (satelliting growth after 24 h, but widespread growth after 48 h of incubation). Cells of G. haemolysans are easily decolorized and resemble those of neisserias, since they occur in pairs with the adjacent sides flattened. G. haemolysans prefers an aerobic growth atmosphere. The esculin hydrolysis test for differentiation of G. haemolysans and R. mucilaginosa in Fig. 2 is performed with esculin agar slants (heart infusion agar containing 0.1% esculin and 0.5% ferric citrate) that are inoculated and incubated at 35°C for up to 7 days. Partial or complete blackening of the agar indicates a positive reaction (5). G. morbillorum cells are Gram positive and arranged in pairs and short chains; individual cells in a given pair may be of unequal sizes. Only a small number of strains of G. bergeri and G. sanguinis have been reported on to date. Information on phenotypic characteristics of

428 n BACTERIOLOGY

these Gemella species can be found in references 18 and 19. The recently described G. asaccharolytica is PYR negative, unlike other Gemella species (20).

Aerococcus The PYR-positive, LAP-negative member of the genus, A. viridans, is characterized by displaying weak or no growth when incubated in an anaerobic atmosphere (134). This trait can be tested by incubating duplicate blood agar plate cultures of the organism in question in anaerobic and aerobic atmospheres and comparing growth after 24 to 48 h. A. viridans forms alpha-hemolytic colonies that could be confused with those of either viridans group streptococci or enterococci. A. sanguinicola is positive in the PYR and LAP tests, while A. urinaehominis is negative in both. The PYRnegative, LAP-positive species, A. urinae and A. christensenii, are differentiated by production of β-glucuronidase (A. christensenii is negative and A. urinae is positive). A. urinae forms small (0.5 mm in diameter after 24 h of incubation), alpha-hemolytic, convex, shiny, transparent colonies on blood agar media. Additional information on the identifying characteristics of A. urinae can be found in reference 107, and a second biotype (esculin hydrolysis positive) of this species is described in reference 24. Additional information on phenotypic traits of the species A. christensenii, A. sanguinicola, and A. urinaehominis can be found in Table 1, Fig. 2, and references 25–28.

Dolosigranulum D. pigrum, the sole species of Dolosigranulum described to date, displays positive PYR and LAP reactions and was initially described as phenotypically similar, though not closely related, to members of the genus Gemella (21). D. pigrum is distinguished from Gemella spp. by its abilities to hydrolyze arginine and to grow in the presence of 6.5% NaCl.

Globicatella and Related Genera (Facklamia, Dolosicoccus, and Ignavigranum) Globicatella and the related genera Facklamia, Dolosicoccus, and Ignavigranum are all, with the exception of the species G. sulfidifaciens, PYR positive. Facklamia and Ignavigranum are also LAP positive and salt tolerant. Globicatella is LAP negative and salt tolerant, while Dolosicoccus is LAP negative and salt intolerant. Dolosicoccus strains are also hippurate hydrolysis negative, which further distinguishes them from strains of Facklamia and Globicatella (hippurate hydrolysis positive). Strains of F. hominis and Ignavigranum may produce urease. Ignavigranum strains may exhibit satelliting behavior. Further details of phenotypic traits of these organisms can be found in references 30–35 and 135.

Helcococcus Colonial morphology (tiny, gray, usually slightly alpha-hemolytic colonies), good growth under anaerobic conditions, and stimulation of growth by addition of 1% horse serum or 0.1% Tween 80 to the medium differentiate H. kunzii from aerococci (36). Isolates of H. kunzii are PYR positive, and most produce an API 20 Strep profile of 4100413. Additional Helcococcus species isolated from humans (H. sueciensis and the proposed “H. pyogenes”) are negative in the PYR test. Detailed phenotypic data on these organisms can be found in references 36–39.

Commercially Available Kits and Automated Methods Based on Phenotypic Traits There have been no comprehensive evaluations of the ability of commercially available products to identify the diverse

and infrequently isolated bacteria described in this chapter. Phenotypic variation among isolates classified in the same species, the relative metabolic inactivity of some organisms, and a relatively small number of strains available for inclusion in databases have challenged the capabilities of these products for accurate identification. Manual methods for performance of some of the basic differentiation tests (e.g., PYR and LAP) are available (e.g., BactiCard Strep; Remel). Commercially available identification kits or systems offering a more comprehensive array of phenotypic tests are improving in their ability to identify many of the organisms discussed in this chapter (26, 135–139). These products include manual methods, e.g., API 20 Strep and RapID Strep (Remel), and automated systems, e.g., VITEK 2 (bioMérieux), MicroScan (Siemens Healthcare Diagnostics, Inc., Deerfield, IL), and Phoenix (BD Diagnostics, Sparks, MD). In the absence of an accurate genus- or species-level identification, these systems will at least provide additional phenotypic information that can be used to augment results of the basic tests mentioned above.

Molecular Methods Sequence-Based Techniques 16S rRNA gene sequencing-based identifications have proven to be more accurate than phenotypic methods for identifying many of the infrequently isolated aerobic catalase-negative, Gram-positive cocci (136, 138). Bosshard and colleagues (136) analyzed 171 clinical strains of the genera Streptococcus, Enterococcus, Abiotrophia, Aerococcus, Granulicatella, and Gemella and observed that more species- or genus-level identifications were achieved based on 16S rRNA gene sequence analysis than based on a commercially available phenotypic method (API 20 Strep), and identifications based on phenotypic traits often disagreed with those determined by 16S rRNA gene sequencing. Woo and coworkers (138) examined strains of Abiotrophia, Granulicatella, Gemella, and Helcococcus in their evaluation of a commercially available 16S rRNA gene sequence-based identification system that analyzes a 527-bp fragment of the 16S rRNA gene (MicroSeq 500; Life Technologies, Foster City, CA). They noted disagreement in identifications obtained with commercially available phenotypic test systems (API 20 Strep and VITEK) and MicroSeq 500 compared with conventional sequencing of the entire 16S rRNA gene. The authors stressed the importance of adequate databases for accurate rRNA gene sequence-based identification. Different regions of the 16S rRNA gene have been used for identification. The first 500 bp is the most applied target for identification. The entire 16S rRNA gene (1.5 kb) has also been sequenced when making more detailed descriptions of taxa or strains (139). 16S rRNA gene sequencing is an effective method for identifying the genera included in this chapter, many of which are difficult to identify with conventional phenotypic methods (139, 140). Accumulating data on the clinical significance of species make it possible to link an accurate species identification to clinically relevant information (139). Other reports on identification of strains of the genera Aerococcus, Granulicatella, and Globicatella support the benefits of using 16S rRNA gene sequencing for these genera (41, 114, 141–143). Broad-range amplification of the 16S rRNA gene directly from clinical samples has made it possible to detect taxa described in this chapter from various clinical samples giving no growth by culture (for instance, G. elegans and A. urinae from cardiac valvular tissue [144, 145]). The sensitivity of

24. Aerobic Catalase-Negative, Gram-Positive Cocci n 429

direct 16S rRNA gene sequencing on valve material is reported to be between 72 and 93% when looking at definite infective endocarditis cases. A major challenge for this methodology is the difficulty in analyzing sequence data when multiple bacterial species are present in the same specimen. Kommedal and coworkers used a set of groupspecific, broad-range primers targeting the 16S rRNA gene followed by DNA sequencing and RipSeq analysis (iSentio AS, Paradis, Norway) and identified Gemella species from abscess material also containing bacterial DNA from other taxa (146). When examining clinical samples, cross-reactivity with human DNA is a major pitfall that can result in mixed electropherograms complicating subsequent sequence analysis and species identification. Kommedal and colleagues employed the principle of dual-priming oligonucleotides to circumvent cross-reactivity even in specimens with a high ratio of human to bacterial DNA (147). Alternative sequencing targets for identification of the organisms discussed in this chapter have also been explored. PCR amplification of a 740-bp rpoB gene fragment followed by sequence analysis was shown to be a suitable molecular approach for the species identification of Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella isolates by Drancourt and colleagues (148). Hung et al. determined the groESL sequences of three species of nutritionally variant streptococci and three Gemella species and developed a multiplex PCR enabling identification of strains of Abiotrophia, Granulicatella, and Gemella at the genus level. To a large extent, intraspecies sequences were well circumscribed, though higher intraspecies heterogeneity was observed in G. haemolysans, with the six isolates examined being separated into two subgroups (149). Tung and coworkers (150) evaluated the feasibility of sequence analysis of the ribosomal 16S-23S intergenic spacer region (ITS) for identification of 24 species of Streptococcus, 1 species of Abiotrophia, 18 species of Enterococcus, and 3 species of Granulicatella. The correct species identification rate by ITS sequence analysis was 98.2%. However, all of the genera except Streptococcus produced more than one PCR amplicon, necessitating agarose gel separation and in some cases cloning to purify the amplicons before sequencing. Tung and colleagues (151) further developed an array-based identification setup based on PCR amplification of the ITS regions and found it to be a useful and reliable alternative to phenotypic identification methods. The approach of having a single molecular platform for strains resembling nonhemolytic streptococci, enterococci, and the taxa considered in this chapter based on ITS analysis has also been examined by Nielsen and colleagues (152). Clinical strains of the genera Aerococcus (n = 37), Abiotrophia (n = 2), Granulicatella (n = 9), Gemella (n = 22), and Leuconostoc (n = 5) were examined. All 75 clinical strains, irrespective of obtained maximum score value, were allocated to the expected species except for 2 strains of Gemella.

MALDI-TOF MS Techniques Matrix-assisted laser desorption ionization–time-of-flight mass spectrometry (MALDI-TOF MS) (see chapter 4) for analysis of bacterial proteins is a promising technique for identification of the bacteria mentioned in this chapter. The MALDI Biotyper (Bruker Daltonics, Billerica, MA) and the VITEK MS (bioMérieux) are two commercially available identification systems employing this technology. Christensen and coworkers (153) examined 51 culture collection strains and 90 well-characterized strains with the Bruker Biotyper system. A mass spectrum profile (MSP)

based on 24 separate determinations was created for each strain and stored in a separate library that could be combined with the standard database (Biotyper version 2.0.43.1), which did not include all the taxa examined. The following genera were represented: Aerococcus (n = 42), Gemella (n = 30), Granulicatella (n = 11), Abiotrophia (n = 3), Lactococcus (n = 9), Globicatella (n = 7), Leuconostoc (n = 9), Rothia (n = 9), Facklamia (n = 7), Vagococcus (n = 2), Helcococcus (n = 4), Alloiococcus (n = 2), Pediococcus (n = 3), Ignavigranum (n = 1), Dolosicoccus (n = 1), and Dolosigranulum (n = 1). The protocol for creating one’s own MSPs in a separate library, though somewhat labor-intensive, makes it possible to extend the database. After extension of the database, all challenge strains were correctly identified to the species level except for a few difficult-to-identify Gemella strains. The obtained mean score values illustrated diversity within the different species and also an effect on mean score values of different conditions for creating MSPs. The MSP dendrograms for Aerococcus, Gemella, Granulicatella, and Abiotrophia convincingly delineated the included species. Thus, for routine use, MALDI-TOF MS was shown to be robust. In a study by Rychert et al., 1,146 strains of Gram-positive cocci were examined with the VITEK MS system (154). Fifteen of these strains belonged to taxa described in this chapter: A. defectiva (n = 2), A. viridans (n = 6), G. haemolysans (n = 3), G. adiacens (n = 1), L. garvieae (n = 1), L. lactis (n = 1), and L. mesenteroides (n = 1). All strains were correctly identified except the L. mesenteroides strain, which gave no identification. Other studies have corroborated the usefulness of MALDI-TOF MS for identification of aerococci (155) and Lactococcus (156, 157) and Leuconostoc (157) species and have investigated using this technique for identification of Weissella and Pediococcus strains (158).

TYPING SYSTEMS Little information exists on typing methods for the genera of infrequently isolated Gram-positive cocci included in this chapter. Typing is not routinely used for characterizing these organisms.

SEROLOGIC TESTS Serologic response to the organisms described in this chapter has not been extensively investigated. No clinically useful tests have been described.

ANTIMICROBIAL SUSCEPTIBILITIES Antimicrobial susceptibility studies on the organisms mentioned in this chapter have generally employed dilution testing methods. Little or no data exist on the utility of disk diffusion or the correlation of Etest results with those of broth or agar dilution methods. Standardized dilution methods and interpretive criteria for observed MICs have been described for only four of the genera (Abiotrophia, Granulicatella, Leuconostoc, and Pediococcus [reference 159 and chapter 74]. The lack of standardized methods and interpretive criteria and the relatively small collections of isolates for some of the genera discussed in this chapter make it difficult to accurately assess antimicrobial susceptibility patterns. With the exception of Leuconostoc, Pediococcus, and Weissella, all of the genera display susceptibility to vancomycin. While many of the genera are susceptible to βlactams and other antimicrobials, observed strain variations suggest that MICs of antimicrobials used for treatment

430 n BACTERIOLOGY

should be determined for individual isolates. When susceptibility testing is requested for isolates for which no guidelines exist, dilution methods may be used to generate MICs that can be reported without interpretation. Since many of the bacteria discussed here are fairly fastidious, investigators have often employed blood-supplemented Mueller-Hinton media and, if necessary for good growth, incubation in a CO2-enriched atmosphere for susceptibility testing. Pyridoxal hydrochloride (final concentration of 0.001%) should also be added to blood-supplemented media for testing strains of Abiotrophia and Granulicatella (159). Details of published susceptibility testing studies for each of the genera appear below. Information on the in vitro antimicrobial susceptibility of L. lactis and L. garvieae strains isolated from humans suggests that L. garvieae isolates are less susceptible to penicillin and cephalothin than are strains of L. lactis. The uniform resistance of L. garvieae versus the uniform susceptibility to clindamycin of the L. lactis strains examined by Elliott and Facklam (129) led them to propose a test for clindamycin susceptibility as an aid in differentiation of these two species. In clinical practice, cases of lactococcal endocarditis have been successfully treated with either penicillin alone or with penicillin and gentamicin (47, 48). Teixeira and colleagues observed that a collection of Vagococcus isolates were all susceptible to ampicillin, cefotaxime, and trimethoprim-sulfamethoxazole. All strains were resistant to clindamycin, lomefloxacin, and ofloxacin. Variable results were observed with other antimicrobial agents (55). The vancomycin-resistant genera Leuconostoc and Pediococcus are penicillin susceptible when MICs are interpreted using criteria adapted from those for Enterococcus spp. (159, 160). They are usually susceptible to chloramphenicol, tetracyclines, and aminoglycosides. Carbapenem and cephalosporin resistance has been noted in some strains of Leuconostoc (159). Huang and colleagues (161) noted MIC ranges of 0.5 to 8 μg/ml for linezolid and 0.06 to 2 μg/ml for daptomycin in 68 strains of Leuconostoc tested and ranges of 1 to 4 μg/ml for linezolid and 0.06 to 0.5 μg/ml for daptomycin in 13 Pediococcus isolates. Iwen and colleagues reported on the successful treatment of a case of P. acidilactici endocarditis with daptomycin (162). Abiotrophia and Granulicatella isolates display a range of penicillin MICs, with authors reporting reduced penicillin susceptibility in 33 to 65% of isolates (163–165). Susceptibility to aminoglycosides is also variable, but no cases of high-level resistance have been reported. A synergistic effect between β-lactam agents and aminoglycosides has been demonstrated for isolates of Abiotrophia, and combination therapy with penicillin and gentamicin is the currently recommended treatment for endocarditis caused by Abiotrophia and Granulicatella. High relapse rates have been reported, even with appropriate therapy (163). Tuohy and colleagues (165) examined a collection of 27 G. adiacens and 12 A. defectiva strains, noting susceptibility of all isolates to clindamycin, rifampin, levofloxacin, ofloxacin, and quinupristindalfopristin. These authors noted that susceptibilities of G. adiacens and A. defectiva, respectively, to other agents tested were as follows: penicillin, 55 and 8%; amoxicillin, 81 and 92%; ceftriaxone, 63 and 83%; and meropenem, 96 and 100% (165). Zheng and coworkers reported high rates of βlactam and macrolide resistance in a collection of pediatric Abiotrophia and Granulicatella isolates (166). A daptomycin MIC range of ≤0.125 to 2 μg/ml was observed for 10 strains of this group of bacteria (167).

A. viridans and G. haemolysans appear to be susceptible to penicillin and display a low level of resistance to aminoglycosides (168, 169). Resistance to tetracycline and macrolides has been described in Gemella isolates (170), as well as a synergistic effect for penicillin and gentamicin (169). Piper and colleagues (167) noted daptomycin MICs of ≤0.125 μg/ml for four strains of G. morbillorum. Buu-Hoi and colleagues (168) noted that while A. viridans seems to be naturally susceptible to macrolides, tetracyclines, and chloramphenicol, resistance to these agents has been observed. A. urinae has been described as susceptible to penicillin, amoxicillin, piperacillin, cefepime, rifampin, and nitrofurantoin but resistant to sulfonamides and netilmicin (107, 171, 172). Humphries and coworkers (173) noted that A. urinae’s resistance to trimethoprim-sulfamethoxazole is test medium dependent. Strains of this organism are susceptible to trimethoprim-sulfamethoxazole when tested on lysed horse blood-containing media that display low levels of thymidine (173). A. sanguinicola isolates display susceptibility to penicillin, amoxicillin, cefotaxime, cefuroxime, erythromycin, chloramphenicol, quinupristin-dalfopristin, rifampin, linezolid, and tetracycline (26). Clinical isolates of D. pigrum studied by LaClaire and Facklam (22) were all susceptible to penicillin, amoxicillin, cefotaxime, cefuroxime, clindamycin, levofloxacin, meropenem, quinupristin-dalfopristin, rifampin, and tetracycline. Variable susceptibility to erythromycin was noted, and 1 of the 27 strains examined was resistant to trimethoprimsulfamethoxazole. The small number of Helcococcus isolates examined displayed susceptibility to penicillin and clindamycin, and most strains were resistant to erythromycin (122, 123). Woo and coworkers described an H. kunzii strain with ermA-mediated erythromycin and clindamycin resistance (126). Strains of Facklamia exhibit variable MICs for a variety of antibiotics (174). A study of 27 strains of Globicatella sanguinis reported susceptibility of all isolates to amoxicillin but varying levels of resistance to other antimicrobials tested (175).

EVALUATION, INTERPRETATION, AND REPORTING OF RESULTS Efforts to identify the Gram-positive cocci included in this chapter should be made only when isolates are considered to be clinically significant (i.e., isolated repeatedly, in pure culture, or from normally sterile sites), since these organisms may also appear in clinical cultures as contaminants or constituents of the normal microbiota. Communication with clinicians should guide the microbiology laboratory in evaluating the significance of these infrequently isolated organisms. The phenotypic tests mentioned in Table 1 and Fig. 1 and 2 facilitate presumptive identification of the infrequently isolated catalase-negative, Gram-positive cocci. More extensive phenotypic testing using commercially available identification systems and molecular methods should be employed for definitive identification. Currently there are susceptibility testing guidelines for only four of the genera mentioned in this chapter. The MICs generated with dilution methods can be reported without interpretation when susceptibility testing is requested for significant isolates for which no guidelines exist. Abiotrophia, Granulicatella, and Gemella species are welldocumented agents of endocarditis. The satelliting behavior of Abiotrophia and Granulicatella and the positive PYR reactions of all three genera are useful for distinguishing them from viridans group streptococci. CLSI guidelines (159)

24. Aerobic Catalase-Negative, Gram-Positive Cocci n

should be employed for susceptibility testing and interpretation of results for Abiotrophia and Granulicatella. The vancomycin-resistant genera Leuconostoc, Pediococcus, and Weissella are infrequent clinical isolates, but they have been described as agents of bacteremia and central nervous system and other infections in compromised hosts. Phenotypic testing for vancomycin resistance (see “Identification” above) is important for identifying these genera and also helps guide antimicrobial therapy. Guidelines for antimicrobial susceptibility testing and interpretation of results are available for Leuconostoc and Pediococcus (159). Among the aerococci, A. urinae is a well-documented urinary tract pathogen and should be reported when isolated in significant amounts as the predominant organism in urine cultures. Phenotypic tests mentioned in this chapter presumptively identify A. urinae, which has been described as susceptible to β-lactam agents and nitrofurantoin (107, 171, 172).

REFERENCES 1. Ludwig W, Schleifer KH, Whitman WB. 2009. Revised road map to the phylum Firmicutes, p 1–17. In Vos P, Garrity GM, Jones D, Krieg NR, Ludwig W, Rainey FA, Schleifer KH, Whitman WB (ed), Bergey’s Manual of Systematic Bacteriology, 2nd ed, vol 3. Springer-Verlag, New York, NY. http://www.bergeys.org/outlines/bergeys_vol_3_ roadmap_outline.pdf. 2. Schleifer KH, Kraus J, Dvorak C, Kilpper-Balz R, Collins MD, Fischer W. 1985. Transfer of Streptococcus lactis and related streptococci to the genus Lactococcus gen. nov. Syst Appl Microbiol 6:183–195. 3. Collins MD, Ash C, Farrow JA, Wallbanks S, Williams AM. 1989. 16S ribosomal ribonucleic acid sequence analyses of lactococci and related taxa. Description of Vagococcus fluvialis gen. nov., sp. nov. J Appl Bacteriol 67:453–460. 4. Wallbanks S, Martinez-Murcia AJ, Fryer JL, Phillips BA, Collins MD. 1990. 16S rRNA sequence determination for members of the genus Carnobacterium and related lactic acid bacteria and description of Vagococcus salmoninarum sp. nov. Int J Syst Bacteriol 40:224–230. 5. Facklam R, Elliott JA. 1995. Identification, classification, and clinical relevance of catalase-negative, gram-positive cocci, excluding the streptococci and enterococci. Clin Microbiol Rev 8:479–495. 6. Collins MD, Lawson PA. 2000. The genus Abiotrophia (Kawamura et al.) is not monophyletic: proposal of Granulicatella gen. nov., Granulicatella adiacens comb. nov., Granulicatella elegans comb. nov. and Granulicatella balaenopterae comb. nov. Int J Syst Evol Microbiol 50:365–369. 7. Kawamura Y, Hou X, Sultana F, Liu S, Yamamoto H, Ezaki T. 1995. Transfer of Streptococcus adjacens and Streptococcus defectivus to Abiotrophia gen. nov. as Abiotrophia adiacens comb. nov. and Abiotrophia defectiva comb. nov., respectively. Int J Syst Bacteriol 45:798–803. 8. Bouvet A, Grimont F, Grimont PA. 1989. Streptococcus defectivus sp. nov. and Streptococcus adjacens sp. nov., nutritionally variant streptococci from human clinical specimens. Int J Syst Bacteriol 39:290–294. 9. Roggenkamp A, Abele-Horn M, Trebesius KH, Tretter U, Autenreith IB, Heesemann J. 1998. Abiotrophia elegans sp. nov., a possible pathogen in patients with culturenegative endocarditis. J Clin Microbiol 36:100–104. 10. Kanamoto T, Sato S, Inoue M. 2000. Genetic heterogeneities and phenotypic characteristics of strains of the genus Abiotrophia and proposal of Abiotrophia para-adiacens sp. nov. J Clin Microbiol 38:492–498. 11. Elliott JA, Facklam RR. 1993. Identification of Leuconostoc spp. by analysis of soluble whole-cell protein patterns. J Clin Microbiol 31:1030–1033.

431

12. Collins MD, Samelis J, Metaxopoulos J, Wallbanks S. 1993. Taxonomic studies on some leuconostoc-like organisms from fermented sausages: description of a new genus Weissella for the Leuconostoc paramesenteroides group of species. J Appl Bacteriol 75:595–603. 13. Barros RR, Carvalho MD, Peralta JM, Facklam RR, Teixeira LM. 2001. Phenotypic and genotypic characterization of Pediococcus strains isolated from human clinical sources. J Clin Microbiol 39:1241–1246. 14. Collins MD, Williams AM, Wallbanks S. 1990. The phylogeny of Aerococcus and Pediococcus as determined by 16S rRNA sequence analysis: description of Tetragenococcus gen. nov. FEMS Microbiol Lett 70:255–262. 15. Ennahar S, Cai Y. 2005. Biochemical and genetic evidence for the transfer of Enterococcus solitarius Collins et al. 1989 to the genus Tetragenococcus as Tetragenococcus solitarius comb. nov. Int J Syst Evol Microbiol 55:589–592. 16. Tunnicliff R. 1917. The cultivation of a micrococcus from blood in pre-eruptive and eruptive stages of measles. JAMA 68:1028–1030. 17. Kilpper-Balz R, Schleifer KH. 1988. Transfer of Streptococcus morbillorum to the genus Gemella as Gemella morbillorum comb. nov. Int J Syst Bacteriol 38:442–443. 18. Collins MD, Hutson RA, Falsen E, Sjoden B, Facklam RR. 1998. Gemella bergeriae sp. nov., isolated from human clinical specimens. J Clin Microbiol 36:1290–1293. 19. Collins MD, Hutson RA, Falsen E, Sjoden B, Facklam RR. 1998. Description of Gemella sanguinis sp. nov., isolated from human clinical specimens. J Clin Microbiol 36:3090–3093. 20. Ulger-Toprak N, Summanen PH, Liu C, Rowlinson MC, Finegold SM. 2010. Gemella asaccharolytica sp. nov., isolated from human clinical specimens. Int J Syst Evol Microbiol 60:1023–1026. 21. Aguirre M, Morrison D, Cookson BD, Gay FW, Collins MD. 1993. Phenotypic and phylogenetic characterization of some Gemella-like organisms from human infections: description of Dolosigranulum pigrum gen. nov., sp. nov. J Appl Bacteriol 75:608–612. 22. Laclaire L, Facklam R. 2000. Antimicrobial susceptibility and clinical sources of Dolosigranulum pigrum cultures. Antimicrob Agents Chemother 44:2001–2003. 23. Aguirre M, Collins MD. 1992. Phylogenetic analysis of some Aerococcus-like organisms from urinary tract infections: description of Aerococcus urinae sp. nov. J Gen Microbiol 138:401–405. 24. Christensen JJ, Whitney AM, Teixeira LM, Steigerwalt AG, Facklam RR, Korner B, Brenner DJ. 1997. Aerococcus urinae: intraspecies genetic and phenotypic relatedness. Int J Syst Bacteriol 47:28–32. 25. Collins MD, Jovita MR, Hutson RA, Ohlen M, Falsen E. 1999. Aerococcus christensenii sp. nov., from the human vagina. Int J Syst Bacteriol 49:1125–1128. 26. Facklam R, Lovgren M, Shewmaker PL, Tyrrell G. 2003. Phenotypic description and antimicrobial susceptibilities of Aerococcus sanguinicola isolates from human clinical samples. J Clin Microbiol 41:2587–2592. 27. Lawson PA, Falsen E, Truberg-Jensen K, Collins MD. 2001. Aerococcus sanguicola sp. nov., isolated from a human clinical source. Int J Syst Evol Microbiol 51:475–479. 28. Lawson PA, Falsen E, Ohlen M, Collins MD. 2001. Aerococcus urinaehominis sp. nov., isolated from human urine. Int J Syst Evol Microbiol 51:683–686. 29. Collins MD, Aguirre M, Facklam RR, Shallcross J, Williams AM. 1992. Globicatella sanguis gen. nov., sp. nov., a new Gram-positive catalase negative bacterium from human sources. J Appl Bacteriol 73:433–437. 30. Collins MD, Falsen E, Lemozy J, Akervall E, Sjoden B, Lawson PA. 1997. Phenotypic and phylogenetic character-

432 n

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

BACTERIOLOGY

ization of some Globicatella-like organisms from human sources: description of Facklamia hominis gen. nov., sp. nov. Int J Syst Bacteriol 47:880–882. Collins MD, Hutson RA, Falsen E, Sjoden B. 1999. Facklamia sourekii sp. nov., isolated from human sources. Int J Syst Bacteriol 49:635–638. Collins MD, Lawson PA, Monasterio R, Falsen E, Sjoden B, Facklam RR. 1998. Facklamia ignava sp. nov., isolated from human clinical specimens. J Clin Microbiol 36:2146– 2148. Lawson PA, Collins MD, Falsen E, Sjoden B, Facklam RR. 1999. Facklamia languida sp. nov., isolated from human clinical specimens. J Clin Microbiol 37:1161–1164. Collins MD, Lawson PA, Monasterio R, Falsen E, Sjoden B, Facklam RR. 1999. Ignavigranum ruoffiae sp. nov., isolated from human clinical specimens. Int J Syst Bacteriol 49:97–101. Collins MD, Rodriguez Jovita M, Hutson RA, Falsen E, Sjoden B, Facklam RR. 1999. Dolosicoccus paucivorans gen. nov., sp. nov., isolated from human blood. Int J Syst Bacteriol 49:1439–1442. Collins MD, Facklam RR, Rodrigues UM, Ruoff KL. 1993. Phylogenetic analysis of some Aerococcus-like organisms from clinical sources: description of Helcococcus kunzii gen. nov., sp. nov. Int J Syst Bacteriol 43:425–429. Collins MD, Falsen E, Brownlee K, Lawson PA. 2004. Helcococcus sueciensis sp. nov., isolated from a human wound. Int J Syst Evol Microbiol 54:1557–1560. Panackal AA, Houze YB, Prentice J, Leopold SS, Cookson BT, Liles WC, Limaye AP. 2004. Prosthetic joint infection due to “Helcococcus pyogenica”. J Clin Microbiol 42:2872–2874. Panackal AA, Houze YB, Prentice J, Leopold SS, Cookson BT, Liles WC, Limaye AP. 2004. Prosthetic joint infection due to “Helcococcus pyogenes”. J Clin Microbiol 42:5966. (Author’s correction.) Kutzer P, Schulze C, Engelhardt A, Wieler LH, Nordhoff M. 2008. Helcococcus ovis, an emerging pathogen in bovine valvular endocarditis. J Clin Microbiol 46:3291–3295. Christensen JJ, Facklam RR. 2001. Granulicatella and Abiotrophia species from human clinical specimens. J Clin Microbiol 39:3520–3523. Reyn A. 1986. Genus Gemella Berger 1960, 253AL, p 1081– 1082. In Sneath PHA, Mair NS, Sharpe ME, Holt JG (ed), Bergey’s Manual of Systematic Bacteriology, vol 2. Williams and Wilkins, Baltimore, MD. Garvie EI. 1986. Genus Leuconostoc van Tieghem 1878, 198AL emend mut. char. Hucker and Pederson 1930, 66AL, p 1071–1075. In Sneath PHA, Mair NS, Sharpe ME, Holt JG (ed), Bergey’s Manual of Systematic Bacteriology, vol 2. Williams and Wilkins, Baltimore, MD. Garvie EI. 1986. Genus Pediococcus Claussen 1903, 68AL, p 1075–1079. In Sneath PHA, Mair NS, Sharpe ME, Holt JG (ed), Bergey’s Manual of Systematic Bacteriology, vol 2. Williams and Wilkins, Baltimore, MD. Elliott JA, Collins MD, Pigott NE, Facklam RR. 1991. Differentiation of Lactococcus lactis and Lactococcus garvieae from humans by comparison of whole-cell protein patterns. J Clin Microbiol 29:2731–2734. Chan JF, Woo PC, Tent JL, Lau SK, Leung SS, Tam FC, Yuen KY. 2011. Primary infective spondylodiscitis caused by Lactococcus garviae and a review of human L. garviae infections. Infection 39:259–264. Mannion PT, Rothburn MM. 1990. Diagnosis of bacterial endocarditis caused by Streptococcus lactis. J Infect 21:317– 318. Pellizzer G, Benedetti P, Biavasco F, Manfrin V, Franzetti M, Scagnelli M, Scarparo C, de Lalla F. 1996. Bacterial endocarditis due to Lactococcus lactis subsp. cremoris: case report. Clin Microbiol Infect 2:230–232.

49. Zechini B, Cipriani P, Papadopoulou S, DiNucci G, Petrucca A, Teggi A. 2006. Endocarditis caused by Lactococcus lactis subsp. lactis in a patient with atrial myxoma: a case report. Diagn Microbiol Infect Dis 56:325–328. 50. Guz G, Colak B, Hizel K, Suyani E, Sindel S. 2006. Peritonitis due to Lactococcus lactis in a CAPD patient. Scand J Infect Dis 38:698–699. 51. Antolin J, Ciguenza R, Saluena I, Vazquez E, Hernandez J, Espinos D. 2004. Liver abscess caused by Lactococcus lactis cremoris: a new pathogen. Scand J Infect Dis 36:490– 491. 52. Guz G, Yegin ZA, Dogan I, Hizel K, Bali M, Sindel S. 2006. Portal vein thrombosis and liver abscess due to Lactococcus lactis. Turk J Gastroenterol 17:144–147. 53. Kim JH, Go J, Cho CR, Kim JI, Lee MS, Park SC. 2013. First report of human acute acalculous cholecystitis caused by the fish pathogen Lactococcus garvieae. J Clin Microbiol 51:712–714. 54. Aubin GG, Bemer P, Guillouzouic A, Cremet L, Touchais S, Fraquet N, Boutoille D, Reynaud A, Lepelletier D, Corvec S. 2011. First report of a hip prosthetic and joint infection caused by Lactococcus garviae in a woman fishmonger. J Clin Microbiol 49:2074–2076. 55. Teixeira LM, Carvalho MG, Merquior VL, Steigerwalt AG, Brenner DJ, Facklam RR. 1997. Phenotypic and genotypic characterization of Vagococcus fluvialis, including strains isolated from human sources. J Clin Microbiol 35:2778–2781. 56. Al-Ahmad A, Pelz K, Schirrmeister JF, Hellwid E, Pukall R. 2008. Characterization of the first oral Vagococcus isolate from a root-filled tooth with periradicular lesions. Curr Microbiol 57:235–238. 57. Al-Tawfiq JA, Kiwan G, Murrar H. 2007. Granulicatella elegans native valve infective endocarditis: case report and review. Diagn Microbiol Infect Dis 57:439–441. 58. Chang SH, Lee CC, Chen SY, Chen IC, Hsieh MR, Chen SC. 2008. Infectious intracranial aneurysms caused by Granulicatella adiacens. Diagn Microbiol Infect Dis 60:201–204. 59. Houpikian P, Raoult D. 2005. Blood culture-negative endocarditis in a reference center: etiologic diagnosis of 348 cases. Medicine 84:162–173. 60. Jeng A, Chen J, Katsivas T. 2005. Prosthetic valve endocarditis from Granulicatella adiacens (nutritionally variant streptococci). J Infect 51:e125–e129. 61. Namdari H, Kintner K, Jackson BA, Namdari S, Hughes JL, Peairs RR, Savage DJ. 1999. Abiotrophia species as a cause of endophthalmitis following cataract extraction. J Clin Microbiol 37:1564–1566. 62. Jeganathan VS, Verma N. 2010. Abiotrophia adiacens-related infectious crystalline keratopathy following penetrating keratoplasty. Br J Ophthalmol 94:139. 63. Cerceo E, Christie JD, Nachamkin I, Lautenbach E. 2004. Central nervous system infections due to Abiotrophia and Granulicatella species: an emerging challenge? Diagn Microbiol Infect Dis 48:161–165. 64. Zenone T, Durand DV. 2004. Brain abscesses caused by Abiotrophia defectiva: complication of immunosuppressive therapy in a patient with connective-tissue disease. Scand J Infect Dis 36:497–499. 65. Atlay M, Akay H, Yildiz E, Duranay M. 2008. A novel agent of peritoneal dialysis-related peritonitis: Granulicatella adiacens. Perit Dial Int 28:96–101. 66. Wilhelm N, Sire S, Le Coustumier A, Loubinoux J, Beljerd M, Bouvet A. 2005. First case of multiple discitis and sacroiliitis due to Abiotrophia defectiva. Eur J Clin Microbiol Infect Dis 24:76–78. 67. Taylor CE, Fang MA. 2006. Septic arthritis caused by Abiotrophia defectiva. Arthritis Rheum 55:976–977.

24. Aerobic Catalase-Negative, Gram-Positive Cocci n 68. del Pozo JL, Garcia-Quetglas E, Hernaez S, Serrera A, Alonso M, Pina L, Leiva J, Azanza JR. 2008. Granulicatella adiacens breast implant-associated infection. Diagn Microbiol Infect Dis 61:58–60. 69. Handwerger S, Horowitz H, Coburn K, Kolokathis A, Wormser GP. 1990. Infection due to Leuconostoc species: six cases and review. Rev Infect Dis 12:602–610. 70. Zaoui A, Brousse C, Bletry O, Augouard LW, Boisaubert B. 2005. Leuconostoc osteomyelitis. Joint Bone Spine 72:79– 81. 71. Dye G, Lewis J, Patterson J, Jorgensen J. 2003. A case of Leuconostoc ventriculitis with resistance to carbapenem antibiotics. Clin Infect Dis 37:869–870. 72. Albanese A, Spanu T, Sali M, Novegno F, D’Inzeo T, Santangelo R, Mangiola A, Anile C, Fadda G. 2006. Molecular identification of Leuconostoc mesenteroides as a cause of brain abscess in an immunocompromised patient. J Clin Microbiol 44:3044–3045. 73. Kumudhan D, Mars S. 2004. Leuconostoc mesenteroids as a cause of post-operative endophthalmitis—a case report. Eye (Lond) 18:1023–1024. 74. Mayeur C, Gratadoux JJ, Bridonneau C, Chegdani F, Larroque B, Kapel N, Corcos O, Thomas M, Joly F. 2013. Faecal D/L lactate ratio is a metabolic signature of microbiota imbalance in patients with short bowel syndrome. PLoS One 8:e54335. doi:doi: 10.1371/journal .pone.0054335. 75. Florescu D, Hill L, Sudan D, Iwen PC. 2008. Leuconostoc bacteremia in pediatric patients with short bowel syndrome: case series and review. Pediatr Infect Dis J 27:1013– 1019. 76. Barton LL, Rider RD, Coen RW. 2001. Bacteremic infection with Pediococcus: vancomycin-resistant opportunist. Pediatrics 107:775–776. 77. Golledge CL, Stingemore N, Aravena M, Joske K. 1990. Septicemia caused by vancomycin-resistant Pediococcus acidilactici. J Clin Microbiol 28:1678–1679. 78. Mastro TD, Spika JS, Lozano P, Appel J, Facklam RR. 1990. Vancomycin-resistant Pediococcus acidilactici: nine cases of bacteremia. J Infect Dis 161:956–960. 79. Sire JM, Donnio PY, Mensard R, Pouedras P, Avril JL. 1992. Septicemia and hepatic abscess caused by Pediococcus acidilactici. Eur J Clin Microbiol Infect Dis 11:623–625. 80. Kumar A, Augustine D, Sudhindran S, Kurian AM, Dinesh KR, Karim S, Philip R. 2011. Weissella confusa: a rare cause of vancomycin-resistant Gram-positive bacteraemia. J Med Microbiol 60:1539–1541. 81. Shin JH, Kim DI, Kim HR, Kim DS, Kook JK, Lee JN. 2007. Severe infective endocarditis of native valves caused by Weissella confusa detected incidentally on echocardiography. J Infect 54:e149–e151. 82. Khan R, Urbin C, Rubin D, Segal-Maurer S. 2004. Subacute endocarditis caused by Gemella haemolysans and a review of the literature. Scand J Infect Dis 36:885–888. 83. Anil M, Ozkalay N, Helvaci M, Agus N, Guler O, Dikerler A, Kanar B. 2007. Meningitis due to Gemella haemolysans in a pediatric case. J Clin Microbiol 45:2337–2339. 84. Lee MR, Lee SO, Kim SY, Yang SM, Seo YH, Cho YK. 2004. Brain abscess due to Gemella haemolysans. J Clin Microbiol 42:2338–2340. 85. Eggelmeijer F, Petit P, Dijkmans BA. 1992. Total knee arthroplasty infection due to Gemella haemolysans. Br J Rheumatol 31:67–69. 86. Kailasanathan A, Anderson D. 2009. Infectious crystalline keratopathy caused by Gemella haemolysans. Cornea 26:643–644. 87. Raman SV, Evans N, Freegard TJ, Cunningham R. Gemella haemolysans acute postoperative endophthalmitis. Br J Ophthalmol 87:1192–1193.

433

88. Ritterband D, Shah M, Kresloff M, Intal M, Shabto U, Seedor J. 2002. Gemella haemolysans keratitis and consecutive endophthalmitis. Am J Ophthalmol 133:268–269. 89. Al-Hujailan G, Lagace-Wiens P. 2007. Mechanical valve endocarditis caused by Gemella morbillorum. J Med Microbiol 56:1689–1691. 90. Gimigliano F, Carletti M, Carducci G, Iodice F, Ballerini L. 2005. Gemella morbillorum endocarditis in a child. Pediatr Infect Dis J 24:190. 91. Valipour A, Koller H, Setinek U, Burghuber OC. 2005. Pleural empyema associated with Gemella morbillorum: report of a case and review of the literature. Scand J Infect Dis 37:378–381. 92. Vasishtha S, Isenberg HD, Sood SK. 1996. Gemella morbillorum as a cause of septic shock. Clin Infect Dis 22:1084– 1086. 93. Benedetti P, Rassu M, Branscombe M, Sefton A, Pellizzer G. 2009. Gemella morbillorum: an underestimated aetiology of central nervous system infection? J Med Microbiol 58:1652–1656. 94. van Dijk M, van Royen BJ, Wuisman PI, Hekker A, van Guldener C. 1999. Trochanter osteomyelitis and ipsilateral arthritis due to Gemella morbillorum. Eur J Clin Microbiol Infect Dis 18:600–602. 95. Roche M, Smyth E. 2005. A case of septic arthritis due to infection with Gemella morbillorum. J Infect 51:e187– e189. 96. Lai CC, Wu CH, Chen JT, Hsueh PR. 2008. Peritoneal dialysis-related peritonitis caused by Gemella morbillorum in a patient with systemic lupus erythematosus receiving steroid therapy. J Microbiol Immunol Infect 41:272–274. 97. Logan LK, Zheng X, Shulman SR. 2008. Gemella bergeriae endocarditis in a boy. Pediatr Infect Dis 27:184–186. 98. Leung DT, Davis EM, Qian Q, Gold HS. 2011. First report of prosthetic joint infection by Gemella sanguinis and associated “pseudosatelliting” phenomenon on culture. J Clin Microbiol 49:3395–3397. 99. Lecuyer H, Audibert J, Bobigny A, Eckert C, JanniereNartey C, Buu-Hoi A, Mainardi JL, Podglajen I. 2007. Dolosigranulum pigrum causing nosocomial pneumonia and septicemia. J Clin Microbiol 45:3474–3475. 100. Hall GS, Gordon S, Schroeder S, Smith K, Anthony K, Procop GW. 2001. Case of synovitis potentially caused by Dolosigranulum pigrum. J Clin Microbiol 39:1202–1203. 101. Lin JC, Hou SJ, Huang LU, Sun JR, Chang WK, Lu JJ. 2006. Acute cholecystitis accompanied by acute pancreatitis potentially caused by Dolosigranulum pigrum. J Clin Microbiol 44:2298–2299. 102. Johnsen BO, Ronning EJ, Onken A, Figved W, Jenum PA. 2011. Dolosigranulum pigrum causing biomaterial-associated arthritis. APMIS 119:85–87. 103. Cattoir V, Kobal A, Legrand P. 2010. Aerococcus urinae and Aerococcus sanguinicola, two frequently misidentified uropathogens. Scand J Infect Dis 42:775–780. 104. Nasoodi A, Ali AG, Gray WJ, Hedderwick DA. 2008. Spondylodiscitis due to Aerococcus viridans. J Med Microbiol 57:532–533. 105. Parker MT, Ball LC. 1976. Streptococci and aerococci associated with systemic infection in man. J Med Microbiol 9:275–302. 106. Grude N, Jenkins A, Tveten Y, Kristiansen BE. 2003. Identification of Aerococcus urinae in urine samples. Clin Microbiol Infect 9:976–979. 107. Christensen JJ, Vibits H, Ursing J, Korner B. 1991. Aerococcus-like organism, a newly recognized potential urinary tract pathogen. J Clin Microbiol 29:1049–1053. 108. Sierra-Hoffman M, Watkins K, Jinadatha C, Fader R, Carpenter JL. 2005. Clinical significance of Aerococcus

434 n

109.

110.

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

121.

122.

123.

124.

125.

126.

BACTERIOLOGY

urinae: a retrospective review. Diagn Microbiol Infect Dis 53:289–292. Delsarte M. 2010. La place des aerococcus en clinique humaine: revue sur une série de 29 cas hospitaliers de 2001 à 2009. Doctoral dissertation. Université Paul Sabatier, Toulouse III, Toulouse, France. Kass M, Toye B, Veinot JP. 2008. Fatal infective endocarditis due to Aerococcus urinae—case report and review of literature. Cardiovasc Pathol 17:410–412. Kristensen B, Nielsen G. 1995. Endocarditis caused by Aerococcus urinae, a newly recognized pathogen. Eur J Clin Microbiol Infect Dis 14:49–51. Santos R, Santos E, Goncalves S, Marques SA, Sequeira J, Abecasis P, Cadete M. 2003. Lymphadenitis caused by Aerococcus urinae infection. Scand J Infect Dis 35:353– 354. Colakoglu S, Turunc T, Taskoparan M, Aliskan H, Kizilkilic E, Demiroglu YZ, Arslan H. 2008. Three cases of serious infection caused by Aerococcus urinae: a patient with spontaneous bacterial peritonitis and two patients with bacteremia. Infection 36:288–290. Ibler K, Truberg Jensen K, Ostergaard C, Wolff Sonksen U, Bruun B, Schonheyder HC, Kemp M, Dargis R, Andresen K, Christensen JJ. 2008. Six cases of Aerococcus sanguinicola infection: clinical relevance and bacterial identification. Scand J Infect Dis 40:761–765. Lau SK, Woo PC, Li NK, Teng JL, Leung DW, Ng KH, Que TL, Yuen KY. 2006. Globicatella bacteraemia identified by 16S ribosomal RNA gene sequencing. J Clin Pathol 59:303–307. Seegmuller I, van der Linden M, Heeg C, Reinert RR. 2007. Globicatella sanguinis is an etiological agent of ventriculoperitoneal shunt-associated meningitis. J Clin Microbiol 45:666–667. Vandamme P, Hommez J, Snauwaert C, Hoste B, Cleenwerck I, Lefebvre K, Vancanneyt M, Swings J, Devriese LA, Haesebrouck F. 2001. Globicatella sulfidifaciens sp. nov., isolated from purulent infections in domestic animals. Int J Syst Evol Microbiol 51:1745–1749. Healy B, Beukenholt RW, Tuthill D, Ribeiro CD. 2005. Facklamia hominis causing chorioamnionitis and puerperal bacteraemia. J Infect 50:353–355. Facklam R. 2002. What happened to the streptococci: overview of taxonomic and nomenclature changes. Clin Microbiol Rev 15:613–630. Haas J, Jernick SL, Scardina RJ, Teruya J, Caliendo AM, Ruoff KL. 1997. Colonization of skin by Helcococcus kunzii. J Clin Microbiol 35:2759–2761. Lemaitre N, Huvent D, Loiez C, Wallet F, Courcol RJ. 2008. Isolation of Helcococcus kunzii from plantar phlegmon in a vascular patient. J Med Microbiol 57:907– 908. Peel MM, Davis JM, Griffin KJ, Freedman DL. 1997. Helcococcus kunzii as sole isolate from an infected sebaceous cyst. J Clin Microbiol 35:328–329. Chagla AH., Borczyk AA, Facklam RR, Lovgren M. 1998. Breast abscess associated with Helcococcus kunzii. J Clin Microbiol 36:2377–2379. Riegel P, Lepargneur JP. 2003. Isolation of Helcococcus kunzii from a post-surgical foot abscess. Int J Med Microbiol 293:437–439. Perez-Jorge C, Cordero J, Marin M, Esteban J. 2012. Prosthetic joint infection caused by Helcococcus kunzii. J Clin Microbiol 50:528–530. Woo PC, Tse H, Wong SS, Tse CW, Fung AM, Tam DM, Lau SK, Yuen K. 2005. Life-threatening invasive Helcococcus kunzii infections in intravenous-drug users and ermA-mediated erythromycin resistance. J Clin Microbiol 43:6205–6208.

127. Ruoff KL, Kuritzkes DR, Wolfson JS, Ferraro MJ. 1988. Vancomycin-resistant Gram-positive bacteria isolated from human sources. J Clin Microbiol 26:2064–2068. 128. Facklam RR, Washington JA II. 1991. Streptococcus and related catalase-negative Gram-positive cocci, p 238–257. In Balows A, Hausler WJ Jr, Herrmann KL, Isenberg HD, Shadomy HJ (ed), Manual of Clinical Microbiology, 5th ed. American Society for Microbiology, Washington, DC. 129. Elliott JA, Facklam RR. 1996. Antimicrobial susceptibilities of Lactococcus lactis and Lactococcus garvieae and a proposed method to discriminate between them. J Clin Microbiol 34:1296–1298. 130. Beighton D, Homer KA, Bouvet A, Storey AR. 1995. Analysis of enzymatic activities for differentiation of two species of nutritionally variant streptococci, Streptococcus defectivus and Streptococcus adjacens. J Clin Microbiol 33:1584–1587. 131. Bouvet A, Villeroy F, Cheng G, Lamesch C, Williamson R, Gutmann L. 1985. Characterization of nutritionally variant streptococci by biochemical tests and penicillinbinding proteins. J Clin Microbiol 22:1030–1034. 132. Davis JM, Peel MM. 1994. Identification of ten clinical isolates of nutritionally variant streptococci by commercial streptococcal identification systems. Aust J Med Sci 15:52–55. 133. Riebel WJ, Washington JA. 1990. Clinical and microbiologic characteristics of pediococci. J Clin Microbiol 28: 1348–1355. 134. Evans JB. 1986. Genus Aerococcus Williams, Hirch and Cowan 1953, 475AL, p 1080. In Sneath PHA, Mair NS, Sharpe ME, Holt JG (ed), Bergey’s Manual of Systematic Bacteriology, vol 2. Williams and Wilkins, Baltimore, MD. 135. LaClaire LL, Facklam RR. 2000. Comparison of three commercial rapid identification systems for the unusual gram-positive cocci Dolosigranulum pigrum, Ignavigranum ruoffiae, and Facklamia species. J Clin Microbiol 38:2037– 2042. 136. Bosshard PP, Abels S, Altwegg M, Bottger EC, Zbinden R. 2004. Comparison of conventional and molecular methods for identification of aerobic catalase-negative gram-positive cocci in the clinical laboratory. J Clin Microbiol 42:2065–2073. 137. Wallet F, Loiez C, Renaux E, Lemaitre N, Courcol RJ. 2005. Performances of VITEK 2 colorimetric cards for identification of gram-positive and gram-negative bacteria. J Clin Microbiol 43:4402–4406. 138. Woo PC, Ng KH, Lau SK, Yip K, Fung AM, Leung K, Tam DM, Que T, Yuen K. 2003. Usefulness of the MicroSeq 500 16S ribosomal DNA-based bacterial identification system for identification of clinically significant bacterial isolates with ambiguous biochemical profiles. J Clin Microbiol 41:1996–2001. 139. Kolbert CP, Rys PN, Hopkins M, Lynch DT, Germer JJ, O’Sullivan CE, Trampuz A, Patel R. 2004. 16S ribosomal DNA sequence analysis for identification of bacteria in a clinical microbiological laboratory, p 361–379. In Persing DH, Tenover FC, Versalovic J, Tang YW, Unger ER, Relman DA, White TJ (ed), Molecular Microbiology: Diagnostic Principles and Practice. ASM Press, Washington, DC. 140. Christensen JJ, Andresen K, Justesen T, Kemp M. 2005. Ribosomal DNA sequencing: experiences from use in the Danish National Reference Laboratory for Identification of Bacteria. APMIS 113:621–628. 141. Christensen JJ, Kilian M, Fussing V, Andresen K, Blom J, Korner B, Steigerwalt AG. 2005. Aerococcus urinae: polyphasic characterization of the species. APMIS 113: 517–525.

24. Aerobic Catalase-Negative, Gram-Positive Cocci n 142. Abdul-Redha RJ, Balslew U, Christensen JJ, Kemp M. 2007. Globicatella sanguinis bacteraemia identified by partial 16S rRNA gene sequencing. Scand J Infect Dis 39:745– 748. 143. Abdul-Redha RJ, Prag J, Sonksen UW, Kemp M, Andresen K, Christensen JJ. 2007. Granulicatella elegans bacteraemia in patients with abdominal infections. Scand J Infect Dis 39:830–833. 144. Maiwald M. 2004. Broad-range PCR for detection and identification of bacteria, p 379–390. In Persing DH, Tenover FC, Versalovic J, Tang YW, Unger ER, Relman DA, White TJ (ed), Molecular Microbiology: Diagnostic Principles and Practice. ASM Press, Washington, DC. 145. Voldstedlund M, Norum Pedersen L, Baandrup U, Klaaborg KE, Fuursted K. 2008. Broad-range PCR and sequencing in routine diagnosis of infective endocarditis. APMIS 116:190–198. 146. Kommedal O, Lekang K, Langeland N, Wiker HG. 2011. Characterization of polybacterial clinical samples using a set of group-specific broad-range primers targeting the 16S rRNA gene followed by DNA sequencing and RipSeq analysis. J Med Microbiol 60:927–936. 147. Kommedal O, Simmon K, Karaca D, Langeland N, Wiker HG. 2012. Dual priming oligonucleotides for broad-range amplification of the bacterial 16S rRNA gene directly from human clinical specimens. J Clin Microbiol 50:1289–1294. 148. Drancourt M, Roux V, Fournier PE, Raoult D. 2004. rpoB gene sequence-based identification of aerobic grampositive cocci of the genera Streptococcus, Enterococcus, Gemella, Abiotrophia, and Granulicatella. J Clin Microbiol 42:497–504. 149. Hung WC, Tseng SP, Chen HJ, Tsai JC, Chang CH, Lee TF, Hsueh PR, Teng LJ. 2010. Use of groESL as a target for identification of Abiotrophia, Granulicatella, and Gemella species. J Clin Microbiol 48:3532–3538. 150. Tung SK, Teng LJ, Vaneechoutte M, Chen HM, Chang TC. 2007. Identification of species of Abiotrophia, Enterococcus, Granulicatella and Streptococcus by sequence analysis of the ribosomal 16S-23S intergenic spacer region. J Med Microbiol 56:504–513. 151. Tung SK, Teng LJ, Vaneechoutte M, Chen HM, Chang TC. 2006. Array-based identification of species of the genera Abiotrophia, Enterococcus, Granulicatella, and Streptococcus. J Clin Microbiol 44:4414–4424. 152. Nielsen XC, Carkaci D, Dargis R, Hannecke L, Justesen US, Kemp M, Hammer M, Christensen JJ. 2013. 16S23S intergenic spacer (ITS) region sequence analysis: applicability and usefulness in identifying genera and species resembling non-hemolytic streptococci. Clin Microbiol 2:130. doi:doi: 10.4172/2327-50733.1000130. 153. Christensen JJ, Dargis R, Hammer M, Justesen US, Nielsen XC, Kemp M, Danish MALDI-TOF MS Study Group. 2012. Matrix-assisted laser desorption ionizationtime of flight mass spectrometry analysis of Gram-positive, catalase-negative cocci not belonging to the Streptococcus or Enterococcus genus and benefits of database extension. J Clin Microbiol 50:1787–1791. 154. Rychert J, Burnham CA, Bythrow M, Garner OB, Ginocchio CC, Jennemann R, Lewinski MA, Manji R, Mochon AB, Procop GW, Richter SS, Sercia L, Westblade LF, Ferraro MJ, Branda JA. 2013. Multicenter evaluation of the Vitek MS matrix-assisted laser desorption ionization-time of flight mass spectrometry system for identification of gram-positive aerobic bacteria. J Clin Microbiol 51:2225–2231. 155. Senneby E, Nilson B, Petersson AC, Rasmussen M. 2013. Matrix-assisted laser desorption ionization-time of

156.

157.

158.

159.

160.

161.

162.

163.

164.

165.

166.

167.

168.

169.

170.

171.

435

flight mass spectrometry is a sensitive and specific method for identification of aerococci. J Clin Microbiol 51:1303– 1304. Tanigawa K, Kawabata H, Watanabe K. 2010. Identification and typing of Lactococcus lactis by matrix-assisted laser desorption ionization-time of flight mass spectrometry. Appl Environ Microbiol 76:4055–4062. De Bruyne K, Slabbinck B, Waegeman W, Vauterin P, De Baets B, Vandamme P. 2011. Bacterial species identification from MALDI-TOF mass spectra through data analysis and machine learning. Syst Appl Microbiol 34:20–29. Albesharat R, Ehrmann MA, Korakli M, Yazaji S, Vogel RF. 2011. Phenotypic and genotypic analyses of lactic acid bacteria in local fermented food, breast milk and faeces of mothers and their babies. Syst Appl Microbiol 34:148–155. Clinical and Laboratory Standards Institute. 2010. Methods for Antimicrobial Dilution and Disk Susceptibility Testing of Infrequently Isolated or Fastidious Bacteria; Approved Guideline—2nd Edition. CLSI Document M45-A. Clinical and Laboratory Standards Institute, Wayne, PA. Swenson JM, Facklam RR, Thornsberry C. 1990. Antimicrobial susceptibility of vancomycin-resistant Leuconostoc, Pediococcus, and Lactobacillus species. Antimicrob Agents Chemother 34:543–549. Huang YT, Liao CH, Teng LJ, Hsueh PR. 2007. Daptomycin susceptibility of unusual gram-positive bacteria: comparison of results obtained by the Etest and the broth microdilution method. Antimicrob Agents Chemother 51:1570–1572. Iwen PC, Mindru C, Kalil AC, Florescu DF. 2011. Pediococcus acidilactici endocarditis successfully treated with daptomycin. J Clin Microbiol 50:1106–1108. Johnson CC, Tunkel AR. 2005. Viridans streptococci and groups C and G streptococci, p 2435–2440. In Mandell GL, Bennett JE, Dolin R (ed), Mandell, Douglas and Bennett’s Principles and Practice of Infectious Diseases, 6th ed. Churchill Livingstone, New York, NY. Liao CH, Teng LJ, Hsueh PR, Chen YC, Huang LM, Chang SC, Ho SW. 2004. Nutritionally variant streptococcal infections at a university hospital in Taiwan: disease emergence and high prevalence of β-lactam and macrolide resistance. Clin Infect Dis 38:452–455. Tuohy MJ, Procop GW, Washington JA. 2000. Antimicrobial susceptibility of Abiotrophia adiacens and Abiotrophia defectiva. Diagn Microbiol Infect Dis 38:189–191. Zheng X, Freeman AF, Villafranca J, Shortridge D, Beyer J, Kabat W, Dembkowski K, Shulman ST. 2004. Antimicrobial susceptibilities of invasive pediatric Abiotrophia and Granulicatella isolates. J Clin Microbiol 42:4323–4326. Piper KE, Steckelberg JM, Patel R. 2005. In vitro activity of daptomycin against clinical isolates of Gram-positive bacteria. J Infect Chemother 11:207–209. Buu-Hoi A, LeBouguenec C, Horaud T. 1989. Genetic basis of antibiotic resistance in Aerococcus viridans. Antimicrob Agents Chemother 33:529–534. Buu-Hoi A, Sapoetra A, Branger C, Acar JF. 1982. Antimicrobial susceptibility of Gemella haemolysans isolated from patients with subacute endocarditis. Eur J Clin Microbiol 1:102–106. Zolezzi PC, Cepero PG, Ruiz J, Laplana LM, Calvo CR, Gomez-Lus R. 2007. Molecular epidemiology of macrolide and tetracycline resistances in commensal Gemella sp. isolates. Antimicrob Agents Chemother 51:1487–1490. Schurr PM, Kasteren ME, Sabbe L, Vos MC, Janssens MM, Buiting AM. 1997. Urinary tract infections with Aerococcus urinae in the south of the Netherlands. Eur J Clin Microbiol Infect Dis 16:871–875.

436 n

BACTERIOLOGY

172. Skov R, Christensen JJ, Korner B, Frimodt-Moeller N, Espersen F. 2001. In vitro antimicrobial susceptibility of Aerococcus urinae to 14 antibiotics, and time-kill curves for penicillin, gentamicin and vancomycin. J Antimicrob Chemother 48:653–658. 173. Humphries RM, Lee C, Hindler JA. 2011. Aerococcus urinae and trimethoprim-sulfamethoxazole. J Clin Microbiol 49:3934–3935.

174. LaClaire L, Facklam R. 2000. Antimicrobial susceptibilities and clinical sources of Facklamia species. Antimicrob Agents Chemother 44:2130–2132. 175. Shewmaker PL, Steigerwalt AG, Shealey L, Weyant R, Facklam RR. 2001. DNA relatedness, phenotypic characteristics, and antimicrobial susceptibilities of Globicatella sanguinis strains. J Clin Microbiol 39:4052–4057.

Gram-Positive Rods

General Approaches to the Identification of Aerobic GramPositive Rods KATHRYN A. BERNARD

25 The purpose of this algorithm for the identification of aerobic Gram-positive rods is to assist the reader in finding the appropriate chapter in this Manual for further information. The algorithm emphasizes that the Gram stain (performed on 24- to 48-h-old colonies from rich media) and macroscopic morphologies are initial key features for the differentiation of aerobic Gram-positive rods. All strains of aerobic Gram-positive rods (except the non-rapidly growing mycobacteria) are initially grown on blood agar plates. Gram-positive organisms demonstrating “regular” rods are those with cells whose longitudinal edges are usually not curved but are parallel. If spore formation is not observed initially, it can be tested for on a nutritionally depleted medium. Catalase activity should be tested with colonies grown on media lacking heme groups. Type of metabolism can be evaluated using oxidative-fermentative media or in cystine Trypticase agar medium. “Irregular” rods are those organisms with cells whose longitudinal edges are curved and not parallel. Slight beta-hemolysis is best observed when cells are incubated in a CO2-enriched atmosphere. Organisms that have yellow- or orange-pigmented colonies are usually composed of irregular rods. Some genera that stain partially acid-fast (e.g., Gordonia and Rhodococcus) may also show a yellow-orange pigment (see chapter 29). Rods exhibiting vegetative substrate filaments may show branched-type hyphae, which either form spores or reproduce by fragmentation. Vegetative substrate filaments might not be present initially (i.e., within 48 h), and so these organisms are prone to being misidentified. For yellow-orange-pigmented genera (e.g., Microbacterium, Curtobacterium, and Leifsonia; see chapter 28), as well as for those rods exhibiting vegetative substrate filaments, chemotaxonomic or molecular genetic methods may be required for definitive identification to the genus level; partially acid-fast bacteria may be identified to the genus level by genetic means. Genera that contain strictly anaerobic Gram-positive rods may also contain species, or strains within a species, that grow reasonably well aerotolerantly or aerobically. This is particularly true for the genera Actinomyces and Actinoba-

culum, some Propionibacterium spp. (see chapter 52), and Clostridium tertium (a strong gas producer) as well as other aerotolerant Clostridium spp. (see chapter 53). Some aerobic Gram-positive cocci, e.g., Leuconostoc spp. (see chapter 24) and Streptococcus mutans (see chapter 22), might initially be misidentified as Gram-positive rods because of their initial Gram stain appearance after growth on plate agar rather than from a broth. Some Gram-positive rods (e.g., Rhodococcus spp. [see chapter 29] or Dermabacter [see chapter 28]) might be misidentified as Gram-positive cocci because of their initial Gram stain appearance. After preliminary examination of the pathogen, the microbiologist should refer to Table 1, where a wide variety of genera of Gram-positive rods are cross-referenced, with respect to relevant chapters in this Manual. Molecular approaches to characterize or subtype pathogens described in this section, as adjuncts to conventional phenotypic or chemotaxonomic assays, are described briefly in each chapter of this Manual, as well as in overview in chapters 6, 10, and 16. Use of 16S rRNA gene sequencing as a means to characterize an isolate has become an established tool in many microbiology laboratories; use of this approach can positively affect patient treatment and facilitate earlier hospital discharge (1). However, interpretation of this work must be done by personnel with a good working knowledge of current approaches for taxonomy and systematics (see chapter 17). Instances exist where 16S rRNA gene sequence analysis cannot be used as the sole characterization method, particularly where two or more validly named taxon groups have >98.7% identity to each other, a cutoff value described as being suggestive of members of the same species (2). In such cases, sequencing of secondary or additional gene targets is recommended (3). Matrix-assisted laser desorption ionization–time-of-flight mass spectrometry is being used increasingly for identification of a wide variety of pathogens (4). The application and efficacy of this technology are outlined for individual taxa within each chapter. Additional reviews describing some of these pathogens are found in references 5–7.

doi:10.1128/9781555817381.ch25

437

438

TABLE 1

Algorithm for identification of Gram-positive rods

Cellular morphology Regular

Irregular

Pigmented

Vegetative Sporesubstrate Catalase Metabolism former filaments

UnusualGram stain features

Acidfast

Partially or weakly acid-fast

Aerial vegetative filaments

Motility

Genus or genera (chapter)

−

−

+

V

O or F

Often large cells

−

−

−

V

−

−

−

+

F

−

−

−

−

+

Bacillus, Paenibacillus, Aneurinibacillus, Virgibacillus, occasionally other genera in family Bacillaceae (26) Listeria (27)

− −

− −

− −

− −

F F

− −

− −

− −

− −

− −

Erysipelothrix (27) Lactobacillus (52)

−, also Y or B-G

−

−

+

O or F

−

−

−

−

Corynebacterium (28)

− −

− −

− −

−, also Y −

− −

− −

+ + + + +

O F F O F

Club shaped; rarely, some unusual forms, e.g., “whip handles” or “bulges” Long slim rods Coccoid Short rods Shorter rods May show branching or short rods

− − − − −

− − − − −

− − − − −

− − − − −

−, also B-G

−

−

+

F

Pleomorphic

−

−

−

−

Turicella (28) Dermabacter (28) Helcobacillus (28) Brevibacterium (28) Actinomyces, Propionibacterium (52 for both) Rothia (21 and 28)

−

−

−

−

O

−

−

−

+

Auritidibacter (28)

−

−

−

−

F

−

−

−

−

Gardnerella (28)

−

−

−

−

F

Coccoidal or rod forms Gram variable, coccoidal Some Actinomyces spp. branching

−

−

−

−

−

−

−

−

F

−

−

−

−

Y, Y-O

+

−

+

F F

−

−

−

V

Arcanobacterium and Trueperella (28 for both), Actinomyces and Actinobaculum (52 for both) Bifidobacterium and genera formerly Bifidobacteriumb (52) Oerskovia (28) Cellulosimicrobium (28)

Pleomorphic, “bifidoforms”

Additional comment Bacillus anthracis, also chapter 14

Motility better observed at 20– 25°C H2S produced in TSI Some strains weakly catalase +

Actinomyces radicidentis, coccoid Some strains catalase − May demonstrate rodcoccus cycle Beta-hemolysis on vaginalis agar Some species with slight betahemolysis

Y, Y-O Y, Y-O

− −

− −

+ +

O O or F

− −

− −

− −

V V

Y, Y-O

−

−

+

O F

−

−

−

+ V+

− −

+ +

− −

+ +

O O

+ +

− −

− −

− −

− −, Y, P-C

+ + or V

− −

+ +

O O

− −

+ + or −

+ −

− −

−

+

−

+

O

−

−

−

+

−, Y

+

−

+

O

−

−

+

−

−

+

−

+

O

−

−

+

−

Dense aggregates of rounded cells

Curtobacterium (28) Microbacterium (28)

Some Microbacterium spp. catalase −

Leifsonia (28) Cellulomonas, Exiguobacterium (28 for both) Mycobacterium (30–32) Segniliparus (29) No growth on MacConkey agar Nocardia (29) Tsukamurella, Dietzia generally not Gordonia, considered acid-fast Rhodococcus, Dietzia, Williamsia (29 for all) Dermatophilus (29) Beta-hemolysis Actinomadura, Streptomyces, Amycolatopsis, Nocardiopsis, occasionally other genera among Pseudonocardiaceae (29 for all) Saccharomonospora, Saccharopolyspora, Thermoactinomyces (29 for all)

Some strains lack aerial filaments

Can grow at 50°C

a Abbreviations and symbols: +, all or nearly all strains positive; −, all or nearly all strains negative; V, feature variable; O, oxidative metabolism; F, fermentative metabolism; TSI, triple sugar iron slant. “Pigment” implies that colonies have pigment other than gray-white or white colony; yellow or yellowish (Y) or yellow-orange (Y-O) pigment is typical, blackish gray (B-G) pigment is occasionally seen, and pinkish coral (P-C) is seen for some Rhodococcus and Williamsia spp. Table excludes extremely infrequently isolated genera described in chapter 28, e.g., Brachybacterium, Knoellia, and Janibacter. b Strains of the genus Alloscardovia in particular can be aerotolerant.

439

440 n

BACTERIOLOGY

REFERENCES 1. Bharadwaj R, Swaminathan S, Salimnia H, Fairfax M, Frey A, Chandrasekar PH. 2012. Clinical impact of the use of 16S rRNA sequencing method for the identification of “difficult-to-identify” bacteria in immunocompromised hosts. Transpl Infect Dis 14:206–212. 2. Stackebrandt E, Ebers J. 2006. Taxonomic parameters revisited: tarnished gold standards. Microbiol Today 33: 152–155. 3. Clinical and Laboratory Standards Institute. 2008. Interpretive Criteria for Identification of Bacteria and Fungi by DNA Target Sequencing; Approved Guideline. CLSI Document MM18-A. Clinical and Laboratory Standards Institute, Wayne, PA.

4. Patel R. 2013. Matrix-assisted laser desorption ionization– time of flight mass spectrometry in clinical microbiology. Clin Infect Dis 57:564–572. 5. Bernard K. 2012. The genus Corynebacterium and other medically relevant coryneform-like bacteria. J Clin Microbiol 50:3152–3158. 6. Savini V, Fazii P, Favaro M, Astolfi D, Polilli E, Pompilio A, Vannucci M, D’Amario C, Di Bonaventura G, Fontana C, D’Antonio D. 2012. Tuberculosis-like pneumonias by the aerobic actinomycetes Rhodococcus, Tsukamurella and Gordonia. Microbes Infect 14: 401–410. 7. Wilson JW. 2012. Nocardiosis: updates and clinical overview. Mayo Clin Proc 87:403–407.

Bacillus and Other Aerobic Endospore-Forming Bacteria* CHRISTINE Y. TURENNE, JAMES W. SNYDER, AND DAVID C. ALEXANDER

26 that is aerobic, mesophilic, catalase positive, and motile by means of peritrichous flagella. Young cultures usually stain Gram positive, whereas older cultures are more likely to be Gram variable or Gram negative. However, the genus is phenotypically diverse such that some species are asporogenous, facultatively anaerobic or strictly anaerobic, and thermophilic or psychrophilic. Many Bacillus species have been renamed and reallocated to new monophyletic and phenotypically coherent taxa, which is consistent with a recent proposal that the designation Bacillus be reserved exclusively for organisms belonging to the Bacillus subtilis and Bacillus cereus clades (2). Even so, the genus Bacillus remains polyphyletic, phenotypically diverse, and large. Frequently encountered species include B. subtilis (the type species), B. licheniformis, B. megaterium, B. pumilus, and the B. cereus group. Also called B. cereus sensu lato, this clade includes B. cereus (sensu stricto), B. anthracis, and B. thuringiensis as well as the less common but phylogenetically related B. mycoides, B. pseudomycoides, B. weihenstephanensis, B. cytotoxicus (143), and B. toyonensis. Other clinically important genera of aerobic endosporeforming organisms are phenotypically similar to the Bacillus, but some differences do exist. Geobacillus species are obligately thermophilic and Geobacillus stearothermophilus (formerly Bacillus stearothermophilus) is commonly employed as a biological indicator for laboratory quality control activities (e.g., for monitoring autoclave efficacy and the sporicidal activity of disinfectants). Lysinibacillus species are distinguished by the composition of the cell wall peptidoglycan, which is rich in lysine and aspartic acid. Lysinibacillus sphaericus (formerly Bacillus sphaericus) occasionally causes opportunistic infections in human hosts but is best known for its insecticidal toxin, which has been commercialized for use in mosquito control programs (3, 4). Paenibacillus and Brevibacillus cultures may stain Gram positive but are more likely to be Gram negative or Gram variable, and sporangia usually have a swollen appearance. Paenibacillus is the type genus of the family Paenibacillaceae and includes species originally defined by 16S rRNA gene sequencing as Bacillus “rRNA group 3” (5). The majority of species are of environmental origin. These include nitrogenfixing organisms associated with plant roots and insect pathogens used for biocontrol applications. Paenibacillus species have been isolated from a variety of clinical sources, including blood, cerebrospinal fluid (CSF), wounds, and urine (6–8).

TAXONOMY Historically, most aerobic endospore-forming bacteria were classified as species of Bacillus. However, the application of phylogenetic methods to prokaryotic systematics has triggered a taxonomic transformation. This includes the valid publication of a new class, Bacilli, within the phylum Firmicutes. Bacilli, as defined by 16S rRNA gene sequence analyses, comprises two orders, Bacillales and Lactobacillales. Currently, the aerobic endospore-forming bacteria are distributed among >60 genera and seven families within the order Bacillales. Minimal standards for describing new taxa of aerobic endospore-forming bacteria have been published (1). These standards acknowledge the role of genetic features but highlight the importance of detailed phenotypic characterization. For any new taxon, it is recommended that descriptions are based on at least five strains and ideally 10 or more. Unfortunately, many new species, and some genera of aerobic endospore-forming bacteria, are represented only by single isolates. The absence of multiple specimens precludes any assessment of intragenus or intraspecies variability. For the clinical microbiologist, the reallocation of taxa and proliferation of new species are of limited practical concern. The vast majority of aerobic endospore-forming bacteria are nonpathogenic, environmental organisms. The medically important and commonly encountered genera, primarily Bacillus, Lysinibacillus, Paenibacillus, and Brevibacillus, are found within the two largest families, Bacillaceae and Paenibacillaceae. The families Alicyclobacillaceae, Planococcaceae, and Sporolactobacillaceae are not known to contain clinically relevant species, and Pasteuria, the only genus in the family Pasteuriaceae, comprises obligate bacterial parasites of invertebrates that have never been grown in axenic culture. In contrast to other Bacillales, organisms in the family Thermoactinomycetaceae appear filamentous and produce mycelia, aerial hyphae, and sporophores. Although some are true aerobic endospore-forming bacteria, for the purposes of this Manual, the Thermoactinomycetaceae are grouped with the aerobic actinomycetes (see chapter 29).

DESCRIPTION OF THE GENERA Bacillus is the type genus of the Bacillaceae. The typical Bacillus isolate is a rod-shaped, endospore-forming organism *This chapter contains information presented by Niall A. Logan, Alex R. Hoffmaster, Sean V. Shadomy, and Kendra E. Stauffer in chapter 24 of the 10th edition of this Manual.

doi:10.1128/9781555817381.ch26

441

442 n

BACTERIOLOGY

Brevibacillus, also in the family Paenibacillaceae, includes species originally defined by 16S rRNA gene sequencing as Bacillus “rRNA group 4” (9). Brevibacillus brevis (the type species), Brevibacillus agri, Brevibacillus centrosporus, and Brevibacillus parabrevis have been isolated from clinical sources (10, 11). Brevibacillus laterosporus is primarily an insect pathogen (12), but human infection has been reported (13). Additional species of aerobic endospore-forming bacteria may be encountered in the microbiology laboratory. Environmental organisms may act as opportunistic pathogens, but most isolates, especially those obtained from nonsterile sites and wound tracts, are likely to be contaminants of unknown clinical significance.

EPIDEMIOLOGY AND TRANSMISSION Aerobic endospore-forming bacteria are ubiquitous in nature. They are found in terrestrial and aquatic habitats of all kinds, ranging from acid to alkaline, nonsaline to highly saline, and hot to cold. Because endospores are resistant to heat, desiccation, radiation, and disinfectants, they persist in places where most other organisms cannot. These exotic and extreme environments include air at high altitude, subterranean waters, glacial ice cores, permafrost, hot springs, and volcanic soils. Dissemination of spores, via wind, dust, and aerosols, contributes to contamination of man-made environments, including hospital operating rooms and industrial clean rooms. Because they contribute to food spoilage and food poisoning, aerobic endospore-forming organisms are unwanted contaminants in food production and preparation environments. However, these bacteria are also essential to the production of certain fermented traditional foods (e.g., natto, cheonggukjang, and gergoush) and the probiotic properties of animal feed and horticultural supplements. Similarly, some of the same species encountered in the clinical laboratory (e.g., B. subtilis, B megaterium, B. licheniformis) are also employed by industry for the commercial production of enzymes (e.g., amylases, proteases, cellulases), antibiotics (e.g., polymyxins, colistins, gramicidins), and pesticides (e.g., Bt toxins). The majority of aerobic endospore-forming bacteria are nonpathogenic saprophytes. Although they are commonly encountered in the microbiology laboratory, only a small minority (∼5%) of isolates are clinically relevant, and most of the clinically significant species are best described as opportunistic human pathogens. Transmission is restricted to ingestion, injection, injury, inhalation, or other contact with material that has been contaminated with spores or vegetative cells. Several environmental species (e.g., B. thuringiensis, Paenibacillus larvae, Paenibacillus lentimorbus, Paenibacillus popilliae, and Pasteuria spp.) are recognized as professional pathogens of insects and other invertebrates, but only B. anthracis has been described as an obligate pathogen of animals. Infection of wild and domesticated mammals occurs through ingestion of spores from contaminated pastures. B. anthracis spores remain viable for many years and are exceedingly difficult to eradicate. Some authorities contend that, in certain soils, self-maintenance can occur (14), and it has recently been demonstrated that soil-dwelling amoebae can support the germination and amplification of B. anthracis spores (15). However, the environmental presence of B. anthracis is generally associated with contamination from animal sources, such as anthrax-infected carcasses. Human cases are almost always due to direct contact with infected animals or animal products. Person-to-person spread has been reported but is rare, as

are cases of anthrax due to laboratory accidents, biological warfare, and bioterror events.

CLINICAL SIGNIFICANCE The majority of aerobic endospore-forming species appear to have little or no pathogenic potential and are rarely associated with disease. The principal exceptions to this are species of the B. cereus group, which includes B. anthracis. However, due to the medical and historical importance of anthrax, B. cereus and B. anthracis are discussed separately below.

Aerobic Endospore-Forming Bacteria (Other Than the B. cereus Group and B. anthracis) Reports of infections due to aerobic endospore-forming bacteria are relatively rare. Cases typically result from contamination of wounds, surgical sites, or medical devices with bacterial spores. Systemic disease most often occurs in patients who are immunocompromised or afflicted with a comorbid condition, but some cases do involve immunocompetent individuals. Mixed infections involving multiple aerobic endospore-forming species have also been reported (16). B. subtilis is the type species of the genus Bacillus. Due to taxonomic changes, some clinical isolates originally identified as B. subtilis have been reallocated to other taxa, but cases involving authentic isolates of B. subtilis do occur. Reports include pneumonia, bacteremia, and septicemia in patients with leukemia or other neoplastic diseases, isolations from surgical wound drainage sites, breast prosthesis and ventriculo-atrial shunt infections, endocarditis following drug abuse (17, 18), and meningitis following a head injury (19). B. subtilis has also been implicated as an agent of food-borne illness and isolated in high numbers (105 to 109 CFU/g) from food. Typical symptoms include vomiting, sometimes accompanied by diarrhea, and reported onset periods have been short (range, 10 min to 14 h; median, 2.5 h) (20). Two cases of severe hepatotoxicity have been attributed to ingestion of B. subtilis-contaminated nutritional supplements (21). A similar case, contamination of an oral probiotic preparation, with an organism ultimately identified as Bacillus clausii, led to a fatal septicemia in an immunocompromised patient. B. clausii was also associated with a case of cholangitis in a renal transplant patient (22). B. licheniformis is another species commonly associated with clinical illness. Its occasional virulence is attributed, at least in part, to the production of lichenilysin, a heatstable and cytotoxic cyclic lipopeptide. B. licheniformis cases include prosthetic valve endocarditis, pacemaker wire infection (23), ventriculitis following the removal of a meningioma, brain abscesses, septicemia following arteriography, bacteremia associated with indwelling central venous catheters, and peritonitis in patients undergoing continuous ambulatory peritoneal dialysis. B. licheniformis contamination of nonsterile cotton wool led to a nosocomial outbreak of bacteremia among patients with blood malignancies (24). Sepsis in immunocompetent patients has also been reported (25). Self-harming behavior was the cause of two unusual cases. In one, B. licheniformis infection followed self-inoculation with drain cleaner (26). In the other, self-inoculation with soil resulted in a polymicrobial infection from which B. licheniformis, B. pumilus, and Paenibacillus polymyxa were isolated (27). B. licheniformis is also an agent of food-borne diarrheal illness, which has led to at least one fatality (28). B. pumilus isolates are known to produce heat-stable, cytotoxic cyclic lipopeptides called pumilacidins. B. pumilus has been implicated in cases of cutaneous, pustule, and rectal

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

fistula infections, bacteremias in immunosuppressed patients (24), a central venous catheter infection in an immunocompetent child, and sepsis in neonates (29), and at least one fatality has been reported (30). Toxigenic strains of B. pumilus have been isolated from cases of food-borne illness and implicated in a rice-associated food poisoning outbreak (31). B. megaterium is occasionally recovered from blood, wounds, and urine. Skin lesions due to B. megaterium can resemble cutaneous anthrax, and phenotypically, the organism can mimic B. anthracis (32, 33). Organisms identified as Bacillus circulans have been isolated from cases of bacteremia in cancer patients, meningitis, CSF shunt infections, endocarditis, wound infections, and peritonitis in a patient undergoing dialysis (34). However, it should be noted that, due to taxonomic changes, some of these isolates might now be classified as species of Paenibacillus. Paenibacillus alvei (formerly Bacillus alvei) has been isolated from cases of meningitis, a prosthetic hip infection in a patient with sickle cell anemia, wound infections, and in association with Clostridium perfringens, a case of gas gangrene. Paenibacillus macerans has been isolated from a wound infection following removal of a malignant melanoma, from a brain abscess following penetrating periorbital injury, and from a catheter-associated infection in a leukemic patient, whereas P. polymyxa has been isolated from patients with bacteremia (27). P. popilliae has been reported from a case of endocarditis, and P. larvae has been reported from infection of a CSF shunt system. Paenibacillus glucanolyticus was involved in cardiac device-related endocarditis (35). Cohnella hongkongensis, a species historically grouped with Paenibacillus (36), was isolated from a case of pseudobacteremia in a boy with neutropenic fever. Lysinibacillus sphaericus has been implicated in a fatal lung pseudotumor, bacteremia, and meningitis (3). Additional clinical cases involve a variety of recently described Bacillus and Paenibacillus species, many of which were proposed on the basis of single isolates of unknown significance from clinical sources: B. idriensis and B. infantis from neonatal sepsis (37); P. konsidensis, P. macerans, P. sanguinis, P. thiaminolyticus, and P. timonensis from blood cultures (6, 8, 30, 38); P. provencensis isolated from CSF (7, 39); P. turicensis from a CSF shunt (40); P. pasadenensis and P. vulneris from wounds (41, 42); and P. urinalis from urine (7). Several Brevibacillus species have also been recovered from clinical sources, including B. centrosporus from bronchoalveolar lavage fluid, B. parabrevis from a breast abscess, and both species from human blood (10). Eye infections due to aerobic endospore-forming bacteria can be quite severe. Many species have been associated with endophthalmitis, including B. circulans, B. licheniformis, P. alvei, and B. laterosporus (43). Bacillus species, including B. megaterium, can cause keratitis. It has also been suggested that B. oleronius, a species found in the gut of Demodex mites, may act as a copathogen in cases of severe or chronic blepharitis (44).

Bacillus cereus Group Bacillus cereus was initially described in an 1887 survey of environmental bacteria (45). The species name is derived from “cera,” the Latin term for wax, and refers to the waxlike morphology of colonies grown on agar media. B. cereus has been isolated from diverse ecological niches and is widely distributed in nature. The original isolate was obtained from the air in a cow shed, and additional strains have been isolated from soil, plant leaves, insects, water, and sewage (46). Psychrotolerant (B. weihenstephanensis), thermotolerant (B. cytotoxicus), crystal-forming (B. thurin-

443

giensis), and morphological (B. mycoides, B. pseudomycoides) variants and a probiotic strain (B. toyonensis) have been classified as distinct species. B. anthracis also belongs to the B. cereus group, but due to the medical and historical importance of anthrax, it will be considered separately. Clinically, B. cereus group organisms are opportunistic pathogens, and cases involving local (e.g., eyes, skin, and wounds) and systemic (e.g., bacteremia, septicemia, meningitis, peritonitis, endocarditis, and respiratory and urinary tract) infections have all been reported (47, 48). Systemic infections are most common in patients who are immunocompromised due to some comorbid condition (e.g., cancer and diabetes). B. cereus spores are resistant to many disinfectants, and postoperative and hospital-acquired infections have been traced to contaminated gloves, gowns, linens, dressings, medical devices (e.g., catheters, shunts and implants, bronchoscopy equipment, and ventilators), and even alcohol-based prep pads used for disinfection purposes (49). A recent nosocomial outbreak, involving 171 patients, was attributed to insufficient air filtration, cleaning, and laundry practices, which failed to eliminate spores that had been introduced to the hospital environment by nearby construction work (50). B. cereus infections secondary to trauma, including cuts and scrapes, surgery, and burns, occur when, during the initial injury, spores from soil, water, or other sources, are introduced to the wound tract. B. cereus contamination of drugs, both legal and illegal, has also been reported (51, 52). Eye injuries and medical procedures (e.g., cataract surgery and intravitreous injections) can result in B. cereus endophthalmitis. Without rapid therapeutic intervention, this serious condition can lead to vision loss and even eye evisceration (43, 53). Keratitis has been associated with contact lens wear, and in at least two cases, B. cereus was cultured from the eyes as well as the contact lens cases of the infected patients (54, 55). However, B. cereus isolates are most commonly associated with food-borne illness. Between 1998 and 2008, B. cereus was suspected or confirmed as the cause in 235 (1.75%) of the 13,405 food-borne outbreaks reported to the Foodborne Diseases Active Surveillance Network (FoodNet) of the Centers for Disease Control and Prevention’s Emerging Infections Program (56). However, that statistic likely underrepresents the true burden of B. cereus disease because these organisms are not always considered in clinical, epidemiological, and laboratory investigations of foodborne illness. Because they survive normal cooking temperatures and many cleaning procedures, B. cereus spores can be widespread in food preparation environments. Food poisoning typically follows the germination of spores and multiplication of toxigenic vegetative cells in improperly stored food. B. cereus group organisms cause two distinct food poisoning syndromes. The emetic type is characterized by nausea and vomiting, with symptoms appearing 1 to 5 h after ingestion of the contaminated food. The diarrheal type is characterized by abdominal pain and diarrhea, and onset is later, 8 to 16 h after food consumption (20, 46). B. cereus has been isolated from diverse foods, but rice dishes, especially Asian-style fried rice, are frequently implicated in outbreaks of emetic type illness, whereas meat dishes are more often associated with diarrheal type illness (56). B. cereus is of increasing concern to the dairy industry. It causes mastitis in goats and dairy cattle, spores persist in barns and processing facilities, and cold-tolerant strains, especially B. weihenstephanensis, can survive both pasteurization and refrigeration (57). Outbreaks of food-borne illness have been linked to milk, cheese, pudding, and other dairy products

444 n

BACTERIOLOGY

(20). Contamination also contributes to food spoilage and shortens the shelf life of dairy products (58).

Bacillus anthracis Robert Koch’s landmark work on anthrax helped establish the germ theory of disease, and an 1877 report on B. anthracis included the first published photomicrographs of any bacteria (59). Anthrax is primarily a disease of wild and domesticated animals that, historically, was a leading cause of mortality among cattle, sheep, goats, and horses. The use of veterinary and human vaccines, improvements in factory hygiene, effective sterilization procedures for imported animal products, and the increased use of synthetic alternatives to animal hides and hair have all contributed to a marked decline in the incidence of disease. Nevertheless, anthrax remains endemic to many countries, particularly those that lack effective vaccination policies. In countries with national vaccination programs, the reduced incidence of anthrax contributes to public ignorance of disease symptoms as well as diminishing veterinary experience. Delays in case recognition can undermine control measures, prolong outbreaks, and lead to the sale and slaughter of affected animals. Because of these challenges, anthrax remains common, especially in agricultural regions of Central and South America, southern and eastern Europe, central and southwestern Asia, sub-Saharan Africa, the United States, and Canada (60). Animals are usually infected through ingestion of B. anthracis spores from contaminated pastures. Direct animalto-animal transmission is rare, although scavengers (e.g., vultures and hyenas) can become infected by feeding on anthrax-infected carcasses (61). Human cases usually involve direct contact with infected animals or products from infected animals. Cases resulting from close contact with infected animals or their carcasses are traditionally classified as nonindustrial, whereas those acquired while processing animal products, such as wool, hair, hides, or bones, are classified as industrial. Although anthrax is not a contagious disease, person-to-person transmission has occasionally been reported, including cases of mother-to-child spread from an infected finger, brother-to-brother spread from an abdominal lesion (62), and nosocomial spread from an umbilical infection (63). Regrettably, B. anthracis has been developed, and effectively deployed, for biological warfare and bioterror purposes. Few reports of laboratory-acquired infections exist, but the accidental release of spores from a military production facility has been implicated as the cause of a major 1979 outbreak in the Ural city of Yekaterinburg (formerly Sverdlovsk), Russia, which claimed at least 66 lives (64). In humans, the major clinical forms of the disease— cutaneous anthrax, gastrointestinal anthrax, inhalational anthrax, and injectional anthrax—are linked to the route of infection (65). Adult males account for the majority of anthrax cases, but disease may be more severe in pregnant women and children (66, 67). Initial symptoms may be nonspecific and mild (e.g., fatigue, malaise, fever, and/or gastrointestinal symptoms), but following lymphohematogenous dissemination from a primary lesion, fulminant disease can rapidly develop. Symptoms can include dyspnea, cyanosis, severe pyrexia, and disorientation, followed by circulatory failure, shock, coma, and death. The bacteria can multiply rapidly in the blood, and depending on the host, reach terminal levels of 107 to 109 CFU/ml over the final few hours. Enhanced clinical and laboratory expertise is a critical component of rapid anthrax diagnosis, and due to the biothreat potential of B. anthracis, there is a need for prospective surveillance and response preparedness. Similarly, case investigations should strive to establish the origin

of any infection so as to differentiate between naturally occurring events and those of malicious origin. Cutaneous anthrax accounts for about 99% of naturally acquired human anthrax cases. In the United States, the incidence of anthrax is extremely low, with 0 to 2 cases reported annually for the last 3 decades (68). The global incidence is difficult to determine and varies between countries depending on control strategies and the strength of health systems. Rates are highest in Africa and central and southern Asia (69) and, in some regions, are increasing due to funding cuts (70). Infection typically occurs when spores are inoculated through a break in the skin, although case reports and animal studies suggest that preexisting lesions are not necessary for infection. Rarely, infection has been attributed to insect bites (71, 72). Following an incubation period of usually 2 to 6 days (range, a few hours to 3 weeks), a small papule appears. Over the next 24 h, this progresses to a ring of vesicles, which subsequently ulcerates to form a blackened eschar, the characteristic lesion of cutaneous anthrax. Once formed, the eschar may become thick and surrounded by extensive edema. Fever, pus, and pain at the site are normally absent, and the presence of such symptoms may be indicative of a secondary bacterial infection. Before the availability of antimicrobial therapy, 10 to 20% of untreated cutaneous anthrax cases were fatal. Today, less than 1% of cases are fatal. Fatalities are mainly due to obstruction of the airways by the edema that accompanies lesions that form on the face or neck but can also occur when cutaneous disease progresses to systemic infection. Eschars take several days to evolve, but even with effective antimicrobial therapy, they may take several weeks to resolve. Gastrointestinal anthrax results from the consumption of anthrax-infected animals, especially meat that is raw or undercooked. Asymptomatic cases can occur, and may not be uncommon, but disease usually presents in one of two forms. The oral, or oropharyngeal, form features lesions on the buccal cavity, tongue, tonsils, or posterior pharyngeal wall. Symptoms include sore throat, dysphagia, and regional lymphadenopathy, followed by severe edema of the neck and chest. The intestinal form can develop anywhere within the gastrointestinal tract, but ulcerations typically appear in the mucosa of the terminal ileum or cecum. Symptoms include nausea, vomiting, anorexia, abdominal pain, fever, mild diarrhea (which can progress to bloody diarrhea), hematemesis, and massive ascites. Owing to the nonspecific nature of the early symptoms, antimicrobial therapy may be initiated late, and mortality rates range widely (73, 74). Inhalational anthrax is a rare but serious form of the disease. During the 20th century, only 18 cases of naturally acquired inhalational anthrax were recorded in the United States. Of these, 16 (89%) were fatal (75). Figures from the United Kingdom show a similar picture. Previously known as pulmonary anthrax, the preferred designation, “inhalational anthrax,” more accurately reflects the fact that active infection occurs in the lymph nodes rather than the lungs themselves. In late 2001, the malicious dissemination of B. anthracis spores in mailed letters was responsible for an outbreak of 22 cases in the United States. Laboratory-confirmed inhalational anthrax accounted for 11 cases, including all 5 of the deaths attributed to the outbreak (76). All of these patients developed severe illness and were hospitalized, but thanks to early recognition and treatment, the case survival rate was 55% (6/11 patients). Analysis revealed a median incubation period of 4 days (range, 4 to 6 days). Clinical presentation included fever or chills, fatigue or malaise, minimal or nonproductive cough, dyspnea, and nausea or vomiting. Some patients experienced chest pain

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n 445

and sweats. All patients had abnormal chest radiographic images, with pleural effusion, infiltrates, or mediastinal widening (77). Injectional anthrax refers to cases associated with injection drug use (78). Although the route of infection is through the skin, symptoms of injectional anthrax differ from those of cutaneous anthrax. Skin around the injection site may be bruised or discolored, but the characteristic papules and eschar are absent. Cases usually present as severe soft tissue infections with significant edema, and progression to septic shock can be rapid (79). Contaminated heroin has been implicated in cases from Norway, Denmark, France, Germany, and the United Kingdom. In Scotland, a large outbreak (47 confirmed, 35 probable, and 37 possible cases) occurred between December 2009 and December 2010 (80). Genotyping methods have been used to analyze B. anthracis isolates associated with cases of injectional anthrax. The European isolates collected between 2000 and 2012 are highly related, share two distinctive single nucleotide polymorphisms, and likely originate from a single source (81, 82). Genotypically, the drug-associated strain is most similar to veterinary isolates from Turkey and not B. anthracis strains from opium (i.e., heroin)-producing countries. Drug traffickers are known to use animal skins for smuggling contraband. As such, it has been hypothesized that the heroin associated with the European cases could have become contaminated with B. anthracis spores if transported in skins from an anthrax-infected animal (80). During the past decade, there have also been several cases of naturally occurring infection. In 2006, a maker of African drums became the first naturally occurring case of inhalational anthrax in the United States since 1976. Infection was traced to imported hides from West Africa which were contaminated with B. anthracis spores. In 2007, two cases of cutaneous anthrax, in a drum maker and a family member, occurred in the United States (83). In 2008, a fatal case of inhalation anthrax occurred in a drum maker in the United Kingdom (84). The handling or playing of goatskin drums contaminated with spores of B. anthracis has been associated with additional cases of cutaneous, inhalational, and gastrointestinal anthrax (83, 85). In rare instances, cases have occurred for which no source has been found. In 2011, a man acquired inhalational anthrax while traveling through four U.S. states, including midwestern states where B. anthracis is enzootic. Despite thorough sampling and the involvement of a dedicated anthrax investigative team, no exposure source was definitively identified (86). A 2012 case from the United Kingdom involved a member of the armed forces who acquired inhalational anthrax despite being vaccinated. An exhaustive inquiry was conducted, but no exposure history to anthrax was identified (87).

Toxins and Other Virulence Factors of the B. cereus Group The pathogenicity of B. cereus group organisms (including B. anthracis) is attributed to the production of numerous exotoxins, enterotoxins, hemolysins, and cytolysins. Ongoing studies aim to define roles for individual toxins, including their distribution, structures, mechanisms of action, and specific contribution to B. cereus disease (88–97). Expression of these toxins is controlled by multiple environmental cues, including nutrient status, oxygen concentration (e.g., redox potential), pH, and cell density. The PlcR-PapR quorum-sensing system regulates expression of at least 45 genes, including those that encode Nhe, HBL, and CytK (see below) (98). Remarkably, this “master” virulence regulator

is inactive in B. anthracis strains, due to a lineage-specific plcR mutation. Two toxins, lethal toxin and edema toxin, are associated with the virulence of B. anthracis. Each of these binary toxins contains two components: a protective antigen (PA) plus an enzymatic factor, lethal factor (LF) for lethal toxin and edema factor (EF) for edema toxin. PA facilitates the delivery of the toxin to target cells, where LF or EF then acts to disrupt cell signaling pathways. The pagC, lef, and cya genes that encode PA, LF, and EF, respectively, are found on a virulence plasmid, pXO1. Loss of pXO1 is associated with attenuation of B. anthracis, but even plasmidcured strains retain some pathogenic potential (88, 89). Conversely, experimental acquisition of pXO1, or natural acquisition of pXO1-like plasmids, can enhance pathogenicity of other B. cereus group organisms. Multiple toxins contribute to the symptoms of B. cereus food poisoning. Ingestion of a heat-stable toxin, cereulide, triggers the vomiting that is characteristic of emetic food poisoning. Cereulide is produced by the plasmid-borne ces gene cluster (90) and is only found in a subset of B. cereus sensu stricto and B. thuringiensis strains (91). Diarrheal symptoms of B. cereus food poisoning are attributed to multiple toxins, including the nonhemolytic enterotoxin (Nhe), hemolysin BL (HBL), and cytotoxin K (CytK; also known as hemolysin IV) (92). The genes responsible for Nhe production are chromosomally encoded (93) and present in most B. cereus group isolates (91). A plasmid-borne variant has been described for B. weihenstephanensis (94). The HBL toxin is also widely distributed. At least two variants have been described (92), but hbl genes appear to be absent from Bacillus cytotoxicus. CytK was first isolated from an enterotoxigenic strain that is now recognized as the type strain of B. cytotoxicus. Despite the absence of cereulide or HBL production, this strain was responsible for an outbreak that included three fatalities. Two variants of the CytK toxin are now recognized: CytK-1 is produced by B. cytotoxicus, whereas CytK-2 is found in other B. cereus group species and is most prevalent among B. cereus sensu stricto and B. thuringiensis. In vitro, the cytotoxic activity of CytK-1 appears to be greater than that of CytK-2 (95). Traditionally, B. thuringiensis strains are defined by their production of parasporal crystals comprised of insecticidal proteins. The pesticidal potential of Bt toxin was recognized in the 1920s and first commercialized in the 1930s. To date, more than 200 variants have been isolated. Bt toxins are used extensively for the control of agricultural pests, and targets include moths, butterflies, beetles, flies, and nematode worms. B. thuringiensis spores can be sprayed onto fields and more recently, crops have been genetically engineered to express the cry and cyt toxin genes (99). Remarkably, these applications do not pose a threat to human health. B. thuringiensis can be isolated from agricultural workers exposed to commercial Bt strains, but reports of illness are rare and no deaths have occurred (100).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS Bacillus Species Other than B. anthracis Clinical specimens for the isolation of Bacillus species other than B. anthracis can be handled using the standard methods (see chapter 18 of this Manual and laboratory biosafety references, such as the CDC’s Biosafety in Microbiological and Biomedical Laboratories) (101, 102). If the transportation time will be less than a few hours, most specimens, including

446 n

BACTERIOLOGY

serum, can be shipped at either room temperature or 2°C to 8°C. If transportation will be overnight or longer, specimens such as stool, sputum, pleural fluid, blood, and material on swabs should be sent at 2°C to 8°C, while fresh tissue and serum samples should be shipped frozen. While blood culture contamination from skin flora may occur despite best practices, rates should not exceed 3% (103). If collecting blood for nucleic acid-based testing (e.g., nucleic acid amplification test [NAAT], PCR), collection tubes containing EDTA or citrate as an anticoagulant are preferable to those containing heparin. Formalin-fixed tissues can be sent at room temperature. In cases of suspected endophthalmitis, vitreous aspirate and biopsy tissue are the optimal specimens. These should be processed as quickly as possible, ideally, at the bedside. Clinical specimens for isolation of Bacillus species other than B. anthracis can be handled safely on the open bench using standard precautions. Efforts should be made to avoid methods that produce aerosols. Any procedures that have the potential to generate aerosols should be performed in a biological safety cabinet. Biosafety level 2 (BSL-2) practices, containment equipment, and facilities are recommended for all activities involving clinical materials and diagnostic quantities of infectious cultures (101). The clinically significant isolates reported to date are of species that grow, and often sporulate, on routine laboratory media at 37°C. Maintenance is simple if spores can be obtained, but it is a mistake to assume that a primary culture or subculture on blood agar will automatically yield spores if it is stored on the bench or in the incubator. It is best to grow the organism for a few days on nutrient agar or Trypticase soy agar containing 5-mg/liter manganese sulfate and refrigerate when microscopy shows that most cells have sporulated. For most species, sporulated cultures on slants of this medium, sealed after incubation, can survive in a refrigerator for years. Alternatively, cultures (preferably sporulated) can be frozen or lyophilized.

Bioterrorism-Related Specimens and the LRN In 1999, the United States Department of Health and Human Services/CDC, in partnership with the Association of Public Health Laboratories and Federal Bureau of Investigation, established a Laboratory Response Network (LRN) to integrate laboratory responses to public health emergencies, including acts of bioterrorism. The LRN links local laboratories (sentinel level) with state (reference level) and federal (national level) laboratories that provide specialized testing and increased biosafety capacity. In all 50 states, there are reference level laboratories able to rapidly detect and confirm the identity of select agents, including B. anthracis. The LRN also provides guidance for testing and transportation of suspicious or challenging specimens. Guidelines for sentinel level laboratories and contact information for state and territorial public health laboratories are available through the CDC (http:// www.bt.cdc.gov/lrn/) and American Society for Microbiology (http://www.asm.org) websites. For general questions, there is a 24-h hotline number [(800) CDC-INFO/ (800) 232-4636] and an e-mail address ([email protected]). LRN consultations may be also requested by calling the CDC Emergency Operations Center [(770) 488-7100]. Although initially limited to the United States, there are now over 150 national and international locations, including laboratories within Canada, Australia, Germany (U.S. military base), Japan (U.S. military base), South Korea (U.S. military base), and the United Kingdom, capable of providing a rapid response to acts of biological terrorism, chemical terrorism, emerging infectious diseases, and other public health threats. Similar programs exist in other countries. In Canada, the National Microbiology Laboratory (NML; Winnipeg, Manitoba, Canada) serves as the federal center and helps coordinate training and testing activities at Canadian LRN (CLRN) sites located in five provinces.

B. anthracis

Specimens from Patients Suspected To Have Anthrax

In many countries, possession of B. anthracis is regulated by legislation. The U.S. Department of Health and Human Services/CDC and the U.S. Department of Agriculture/APHIS define B. anthracis as a tier I, category A select agent. The Canadian Human Pathogen and Toxins Act (http://loislaws.justice.gc.ca/eng/acts/H-5.67/FullText.html) defines B. anthracis and anthrax toxin as schedule 3 and schedule 1 agents, respectively. Human infectious doses have not been established, but using data based largely on nonhuman primate studies, the U.S. Department of Defense estimates that a 50% lethal dose for humans is 8,000 to 10,000 B. anthracis spores. When collecting clinical specimens for suspected anthrax, appropriate personal protective equipment should be used, including disposable gloves, disposable apron or overalls, and boots which can be disinfected after use. The use of a face shield and/or a respirator should be considered, especially for dusty samples that might contain many spores. Full details of personal protective equipment and of disinfection and decontamination are given in annexes 1 and 3 of the WHO Anthrax in Humans and Animals guidelines (69). It should be noted that waterless rubs containing ethanol are not effective at removing endospores. Hand washing with soap and water or with chlorhexidine gluconate, and the use of hypochloritereleasing towels, may reduce endospore contamination of the skin. Preexposure vaccination recommendations from the Advisory Committee on Immunization Practices (ACIP) are summarized below (104).

The CDC has published guidelines for clinical evaluation of persons with possible anthrax (http://www.cdc.gov/mmwr/ preview/mmwrhtml/mm5043a1.htm). In all cases, specimens from potential sources of infection (e.g., animal hides, hair, and carcasses, etc.) should be sought in addition to patient specimens. Due to the hazardous nature of B. anthracis, it is recommended that sentinel level laboratories refer testing to a LRN reference level, or higher, laboratory. The preferred diagnostic specimen depends on the form of the disease (69). Whenever possible, patient specimens should be collected prior to the initiation of antimicrobial therapy. If cutaneous anthrax is suspected, the edge of an eschar should be lifted and two specimens of vesicular fluid collected by rotating swabs beneath it. One swab is used for Gram stain and culture and the other for NAAT. Immunohistochemical analyses are rarely performed any more, but if such testing is available, a full-thickness punch biopsy specimen from a papule or vesicle lesion that includes adjacent skin should be taken and fixed in 10% buffered formalin. If vesicle and eschar are present, biopsy specimens should be taken from both (105). Inhalational anthrax will be suspected only if the patient’s history suggests it. Chest radiographs and chest computed tomography scans are recommended. Blood should be collected for culture. Serology is useful for the diagnosis of cases where culture fails owing to previous treatment. Acute-phase serum (obtained within 7 days of onset) and convalescent-phase samples (obtained

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

2 to 3 weeks later) should be collected for serologic testing at a federal center (e.g., CDC, NML). Pleural fluid, if present, should be obtained for Gram stain, culture, and PCR. Pleural and/or bronchial biopsy samples can also be tested using immunohistochemical methods. Gastrointestinal anthrax will only be suspected if an adequate history of the patient is known. Specimens from oral lesions may be collected in the same way as those for cutaneous disease and processed for Gram stain, culture, and NAAT. Blood should be obtained for culture, whereas acute- and convalescent-phase serum samples for serological testing should be collected 2 to 4 weeks apart. A stool or rectal swab and, if present, ascites fluid should be collected for Gram stain, culture, and NAAT. Any hemorrhagic fluid from the nose, mouth, or anus should be cultured. Anthrax meningitis may occur on its own or, more frequently, subsequent to other forms of the disease. CSF and blood specimens should be collected for culture, Gram stain, and NAAT. Specimens should also be subjected to antigen detection testing, if available. Postmortem samples are also of diagnostic value. A characteristic of anthrax is nonclotting blood at death. As such, blood for culture, stains, and NAAT may be collected by venipuncture. If these are positive, no further specimens are needed. If negative, additional postmortem specimens should be collected. For example, in cases of suspected gastrointestinal anthrax, peritoneal or ascites fluid, spleen, and/ or mesenteric lymph node specimens may be informative. Care should be taken to use techniques that avoid spillage of fluids. Additional information regarding the collection and transport of specimens that may contain B. anthracis is described in chapter 14 of this Manual, the LRN Sentinel Level Clinical Laboratory Protocols website (http://www.asm.org/ index.php/guidelines/sentinel-guidelines), the Canadian Transportation of Dangerous Goods regulations website (http://www.tc.gc.ca/eng/tdg/clear-menu-497.htm), and the Clinical Microbiology Procedures Handbook (106). In the United States, shipment of B. anthracis requires completion of the APHIS/CDC form 2 (Request to Transfer Select Agents an d T ox in s; ht tp :/ /w ww .s el ec ta ge nt s. go v/ TransferForm.html) plus prior approval from either the CDC or APHIS (see http://www.selectagents.gov/ for additional information about regulation of select agents and toxins in the United States). Nonregistered clinical laboratories, sentinel level, must adhere to current packing and shipping guidelines (LRN Packing and Shipping Guidelines; www.asm.org) for mailing or transferring infectious substances (107). Leakproof containers must be used, then placed in secondary containers for “double-bagging,” and sealed in secure, outer containers for carriage.

Specimens from Animals Suspected To Have Anthrax Anthrax should be considered as the possible cause of death in herbivorous animals that have died suddenly and unexpectedly, especially if hemorrhage from the nose, mouth, or anus has occurred, or if death has taken place at a site with any history of anthrax (e.g., even from several decades before). Specimen collection is influenced by the type of animal involved and the time since death (61, 69). With animals that have died of anthrax, B. anthracis is rapidly destroyed by putrefaction in the intact carcass. However, sporulation will still occur in some tissues, so a carcass generally yields positive cultures. For fresh carcasses (1 to 2 days old), appropriate specimens include peripheral blood and tissue biopsy specimens, such as snips from the tip of an ear. For direct demonstration of B. anthracis using Gram,

447

Giemsa, or polychrome methylene blue (M’Fadyean) staining, hemorrhagic exudate is preferred to aspirated blood or tissue specimens. In pigs, the enormous terminal bacteremia seen in herbivores may not develop, so blood may not be as informative as other bodily fluids. When cervical edema is present, smears and cultures should be made of fluid aspirated from the enlarged mandibular and suprapharyngeal lymph nodes. Intestinal anthrax of pigs may be obvious only at necropsy, but B. anthracis is usually visible in stained smears made from mesenteric lymph nodes. Because B. anthracis competes poorly with putrefactive organisms, it may not be visible in smears prepared from older carcasses (2 to 3 days old). Instead, sections of tissue, or any blood-stained material, should be collected for culture. If the animal has been opened, spleen or lymph node specimens should also be taken. With putrefied and very old carcasses, swabs of the nostrils, nasal turbinates, and eye sockets are likely to yield B. anthracis, but the best specimens may be samples of contaminated soil taken from beneath the head and tail.

B. anthracis Vaccines and Vaccination Anthrax Vaccine Adsorbed (AVA, also called BioThrax; see http://www.fda.gov/BiologicsBloodVaccines/Vaccines/ ucm061751.htm) is the current human vaccine in the United States. It is a cell-free filtrate (formalin treated) in an aluminum hydroxide-adsorbed gel prepared from a noncapsulated, nonproteolytic derivative of strain V770NP1-R grown under microaerobic conditions. The FDA has approved a new schedule for preexposure immunization with AVA such that the five-dose primary schedule has been replaced by a three-dose primary schedule (intramuscular injections at 0, 1, and 6 months) with boosters at 12 and 18 months (http://www.fda.gov/BiologicsBloodVaccines/ Vaccines/ApprovedProducts/ucm304758.htm). To maintain immunity, an annual booster injection is recommended. Anthrax Vaccine Precipitated (AVP) is the current human vaccine in the United Kingdom. It is an alum-precipitated cell-free filtrate of the Sterne strain (34F2) cultured under static batch conditions, with activated charcoal, to increase PA production. Both AVA and AVP contain PA as well as trace amounts of LF, EF, and cell wall proteins. In 2010, the updated ACIP recommendations for pre-event vaccination and postexposure prophylaxis were published (104). In addition to previously approved recommendations for occupational and laboratory populations, the updated recommendations included new language to address emergency responders. The ACIP recommends routine preexposure vaccination with AVA for persons engaged in work (i) with high concentrations or pure cultures of B. anthracis spores, (ii) with environmental samples associated with anthrax investigations, or (iii) in spore-contaminated areas or other settings with exposure to aerosolized B. anthracis spores. Immunization is not routinely recommended for emergency and other responders but may be offered on a voluntary basis as part of a comprehensive occupational health and safety program to persons who perform site investigations, respond to suspicious substance reports (e.g., white powder incidents), or perform other activities that might lead to exposure to aerosolized B. anthracis spores. Because workers in the general diagnostic laboratories, using standard BSL2 practices in the routine processing of clinical or environmental specimens, are not at increased risk for exposure to B. anthracis spores, immunization with AVA is not recommended (104). New anthrax vaccines, including secondand third-generation products, as well as several human monoclonal and polyclonal antibody products, are currently in their early development and testing (108).

448 n

BACTERIOLOGY

FIGURE 1 (a) Gram stain of B. anthracis, associated with a bioterrorism attack, showing Grampositive rods in peripheral blood buffy coat following admission of patient. Bar, 3 μm. (Courtesy of H. Masur.) (b) Spore-stained preparation of Bacillus cereus sporangia, viewed by bright-field microscopy. Spores are stained green, and vegetative cells are counterstained red. Bar, 2 μm. (Photograph kindly provided by M. Rodríguez-Díaz.) doi:10.1128/9781555817381.ch26.f1

DIRECT EXAMINATION Microscopic examination of Gram-stained smears remains a primary tool of the microbiology laboratory (Fig. 1a). Clinically significant aerobic, Gram-positive bacilli include various species of Corynebacterium, Listeria, Lactobacillus, and Nocardia, among others, but Bacillus species are readily distinguished by their microscopic and phenotypic morphology. Aerobic endospore-forming organisms do not always stain Gram positive, but the presence of unstained areas within the cell may be indicative of spores. Phase-contrast microscopy (at a magnification of ×1,000) allows spores to be distinguished from other kinds of inclusions, such as polyhydroxybutyrate (PHB) granules. Spores are larger, more phase bright, and more regular in shape, size, and position. Sporangial appearance is also valuable for specieslevel identification. Although less convenient, a Gramstained smear can be stripped of oil with acetone-alcohol, washed, and then stained for spores. Based on the SchaefferFulton endospore stain (109), a heat-fixed smear is flooded with 10% aqueous malachite green for up to 45 min (without heating), then washed and counterstained with 0.5% aqueous safranin for 30 s. When visualized at 1,000× magnification, the cells are pink-red and spores, if present, are green (Fig. 1b). The polychrome methylene blue (M’Fadyean) staining test allows visualization of the capsule, a characteristic feature of B. anthracis (Fig. 2a). Direct examination of blood smears can reveal capsulated rods or, if blood was collected after treatment was initiated, capsule “ghosts.” Notably, capsule visualization is not recommended in the LRN sentinel level clinical laboratory guidelines, and staining of B. anthracis should only be performed by reference level laboratories. For putrefied specimens, Giemsa stain may be more effective than the M’Fadyean test. The genes required for capsule biosynthesis (capB, -C, -A) are located on the pXO2 virulence plasmid (110). Loss of pXO2 gives rise to nonencapsulated B. anthracis strains which are frequently mistaken for B. cereus. Similarly, species other than B. anthracis occasionally acquire genes for capsule biosynthesis, which may confound staining methods as well as NAAT assays that target capBCA (see below).

Molecular and antigen-based methods for direct examination of specimens exist, but most have been developed for the detection of B. cereus or B. anthracis. The symptoms associated with B. cereus food poisoning are attributed to bacterial toxins. The enterotoxin complex responsible for the diarrheal type of B. cereus food poisoning has been increasingly well characterized (92). Tissue culture-based assays have been developed, and two commercial kits are available for the detection of enterotoxin in foods and feces. The Oxoid BCET-RPLA (Oxoid Ltd., Basingstoke, United Kingdom) and TECRA VIA (TECRA Diagnostics, Roseville, New South Wales, Australia) detect different antigens, and there is some controversy about their reliabilities. Bioassays using human (HEp-2, HepG2) and mouse (Hepa-1) cell lines, boar spermatozoa, and bacterial cultures have been developed for the nonspecific detection of the B. cereus emetic toxin in food extracts and culture filtrates (111). Specific detection of cereulide requires high-performance liquid chromatography–mass spectrometry methods (112). Toxin profiling can also be performed using real-time PCR assays that target toxin biosynthesis genes (113). Because of its pathogenicity and biothreat potential, it is critical for laboratory staff to be able to distinguish B. anthracis from morphologically similar bacteria. Whenever large, nonmotile, Gram-positive rods are observed, and especially if large numbers of these bacilli are observed in a patient’s blood at death, then B. anthracis should be suspected. The LRN provides protocols for ruling out B. anthracis and referring out potential isolates. Several methods, including a B. anthracis-specific LRN PCR assay (114) and serology (115), were effectively used during investigation of the 2001 bioterrorism-associated outbreak as well as for confirmation of more recent cases associated with drum making (83, 105). The most widely used and efficacious detection method in the U.S. public health system is the LRN PCR (114). This test targets several distinct loci on the B. anthracis chromosome and pXO1 and pXO2 virulence plasmids. The use of multiple loci increases specificity and allows for the detection of avirulent or noncapsulated B. anthracis strains (e.g., lacking pXO1 or pXO2), that might otherwise be mistaken for some other organism (e.g., B. cereus). At the

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

FIGURE 2 Photomicrographs of endospore-forming bacteria viewed by bright-field microscopy (a) and phase-contrast microscopy (b to l). Bars, 2 μm. (a) B. anthracis, M’Fadyean stain showing capsulate rods in guinea pig blood smear; (b) B. cereus, broad cells with ellipsoidal, subterminal spores, not swelling the sporangia; (c) B. thuringiensis, broad cells with ellipsoidal, subterminal spores, not swelling the sporangia, and showing parasporal crystals of insecticidal toxin (arrows); (d) B. megaterium, broad cells with ellipsoidal and spherical, subterminal and terminal spores, not swelling the sporangia, and showing PHB inclusions (arrows); (e) B. subtilis, ellipsoidal, central and subterminal spores, not swelling the sporangia; (f) B. pumilus, slender cells with cylindrical, subterminal spores, not swelling the sporangia; (g) B. circulans, ellipsoidal, subterminal spores, swelling the sporangia; (h) Lysinibacillus sphaericus, spherical, terminal spores, swelling the sporangia; (i) Brevibacillus brevis, ellipsoidal, subterminal spores, one swelling its sporangium slightly; (j) Brevibacillus laterosporus, ellipsoidal, central spores with thickened rims on one side (arrow), swelling the sporangia; (k) Paenibacillus polymyxa, ellipsoidal, paracentral to subterminal spores, swelling the sporangia slightly; (l) Paenibacillus alvei, cells with tapered ends, ellipsoidal, paracentral to subterminal spores, not swelling the sporangium. doi:10.1128/9781555817381.ch26.f2

449

450 n

BACTERIOLOGY

CDC, a positive PCR result on any clinical specimen from a patient collected from a normally sterile site (such as blood or CSF) or a lesion of other affected tissue (e.g., skin, pulmonary, reticuloendothelial, or gastrointestinal) is regarded as a supportive or presumptive diagnostic test. It is considered sufficient to provide a probable diagnosis but is not confirmatory in itself. The principal reason for such stringent guidelines on the use of NAAT approaches and the value of their results towards providing a confirmatory diagnosis is based on the possibility that environmental contamination of a non-anthrax-related lesion could result in a positive result. This is especially the case with the use of some previously published PCR primers for capsule and chromosomal genes that can produce false positives with reactions to soil microbiota. This is quite similar to the recommendations that are included in the 2008 WHO guidelines (69), in which PCR can be used for identification of an isolate but is not recommended for testing of specimens. A two-component direct fluorescent-antibody assay was used to identify encapsulated vegetative cells of B. anthracis (116), but NAAT-based detection methods are now preferred. An antibody specific for one of the B. anthracis S-layer proteins is the basis of RedLine Alert (Tetracore, Inc., Gaithersburg, MD), a rapid test that can provide a result within 15 min. This assay has been approved by the FDA for use on nonhemolytic Bacillus species colonies cultured on sheep blood agar plates. Manufacturer’s data suggest that the test was 98.6% sensitive when tested on 145 B. anthracis isolates and 45 nonhemolytic, non-B. anthracis isolates. However, such identification of B. anthracis is only considered presumptive, and this test should not be used as a stand-alone test. The Bacillus anthracis immunochromatographic field assay, developed by the U.S. Naval Medical Research Center (Silver Spring, MD), is an immunochromatographic assay for detection of PA in blood samples and tissue exudates. The highly sensitive and specific assay has been used to detect B. anthracis in animals, even several days after death (117). Numerous reports describe the use of alternative technologies, such as flow cytometry, differential pulse voltammetry, fluorescence resonance energy transfer, surface-enhanced Raman scattering, locked nucleic acid probes, and atomic force microscopy for the detection of B. anthracis (106, 118, 119). Because anthrax toxins are highly expressed during infection, methods for direct detection of PA and LF in specimens (i.e., instead of bacilli) have also been explored. These include immunoassays that target PA (120) as well as bioassays that exploit the metalloprotease activity of LF and the adenylyl cyclase activity of EF (121). Direct detection of spores in environmental samples has been achieved using diagnostic electron microscopy (122), whereas a technique developed by the Norwegian Defence Research Establishment uses matrix-assisted laser desorption ionization–time of flight (MALDI-TOF) mass spectrometry (MS) for examination of suspicious powders (123). The MALDITOF method can accurately distinguish spores from common hoax materials and differentiate B. anthracis from other Bacillus species. Inactivation of BSL-3 organisms, an essential step prior to performing MALDI-TOF (MS) in a BSL-2 space, has been successful for B. anthracis using trifluoroacetic acid (124) and gamma irradiation (125). A genetically engineered B. anthracis-specific bioluminescent reporter phage was capable of detecting B. anthracis, in clinically relevant concentrations, from simulated blood

specimens in as little as 5 h (126) and could also be used for rapid susceptibility testing. These, and other novel approaches, may allow for detection of B. anthracis infection earlier in the course of the disease and thus allow for more successful treatment.

ISOLATION PROCEDURES Isolation of Aerobic Endospore-Forming Bacteria Specimens from patients should be inoculated onto plates of blood agar according to standard methods. Though not commonly used in clinical laboratories, heat treatment (e.g., 70°C for 30 min or 80°C for 10 min) may be used as an enrichment procedure for the isolation of aerobic endospore-forming bacteria. The heat-resistant spores survive temperatures that kill other organisms (e.g., non-sporeforming contaminants), and spores may also be heat shocked into subsequent germination. In general, enrichment procedures are not appropriate for fresh clinical specimens because spores are usually sparse or absent, and heat treatment will kill any vegetative cells. However, for episodes of food poisoning where B. cereus is suspected, nutrient or tryptic soy broth with polymyxin (100,000 U/liter) may be added to a heat-treated stool specimen. For the majority of aerobic endospore-forming species, selective media are not available. However, such media have been designed for the isolation, identification, and enumeration of B. cereus group organisms. These exploit the phenotypic features of the organism, including the positive egg yolk reaction (i.e., phospholipase C activity) and the negative acid-from-mannitol reaction. Addition of pyruvate and polymyxin can enhance the selectivity of these media. Effective formulations include BCM (Bacillus cereus medium; LabM, Heywood, United Kingdom), PEMBA (polymyxin B, egg yolk, mannitol, bromthymol blue agar; Oxoid), and MEYP (mannitol, egg yolk, polymyxin B agar; Oxoid), which is also called MYP (Difco, BD, Franklin Lakes, NJ). Newer formulations, including Bacillus cereus group plating medium (Biosynth Chemistry and Biology, Staad, Switzerland, also available as Cereus-Ident-Agar from Heipha, Eppelheim, Germany) and Bacillus cereus/Bacillus thuringiensis chromogenic plating medium (R&F Laboratories, Downers Grove, IL), use specific chromogenic substrates, rather than natural egg yolk, to reveal phospholipase C activity. Selective media for B. anthracis include PLET agar (polymyxin-lysozyme EDTA-thallous acetate) and the differential/selective chromogenic medium, R&F Anthracis chromogenic agar (R&F Laboratories). Independent studies indicate that both formulations are effective for isolation of B. anthracis, but PLET agar may be more sensitive and selective.

Isolation of B. anthracis Provided that standard good laboratory practice is observed, isolation and procedures to rule out B. anthracis can be performed safely in the routine clinical microbiology laboratory (e.g., by sentinel level laboratories). In the case of a suspect identification of B. anthracis, the isolate must be referred to an LRN reference center equipped for confirmatory identification of B. anthracis. Due to the infectious nature of this pathogen, all manipulations of specimens should be handled in a BSL-2 facility using BSL-3 precautions (see chapter 14 of this Manual and Biosafety in Microbiological and Biomedical Laboratories) (101). BSL-3 facilities are recommended for laboratory personnel doing routine processing of clinical or environmental specimens, but pre-

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

exposure vaccination is not (104). When working with pure cultures of B. anthracis, the primary hazards to laboratory personnel are direct and indirect contact of broken skin with cultures and contaminated laboratory surfaces, accidental parenteral inoculation, and, rarely, exposure to infectious clinical specimens. Laboratories that frequently centrifuge B. anthracis suspensions should use an aerosol-tight rotor that can be repeatedly autoclaved (101). In the United States, occupational exposure or release of the organism outside the primary barriers of the biocontainment area (e.g., on the open bench) requires the completion of APHIS/ CDC form 3 (“Information Document for Report of Theft, Loss, or Release of Select Agents and Toxins”; http:// www.selectagents.gov/TLRForm.html). Possession of B. anthracis requires registration of a laboratory within the United States with either the CDC or APHIS. When B. anthracis is identified by a laboratory, the identification of this agent must be reported to the CDC or APHIS immediately and an APHIS/CDC form 4 (“Report of the Identification of a Select Agent or Toxin”; http:// www.selectagents.gov/CDForm.html) submitted within 7 days. Other authorities should be notified as required by federal, state, or local laws. When B. anthracis is isolated in an unregistered laboratory (e.g., diagnostic sentinel level clinical laboratory), the organism must either be destroyed on-site by a recognized sterilization or inactivation process or be transferred to a registered laboratory within 7 days following notification of the isolate having been confirmed as B. anthracis. In Canada, all materials containing or suspected to contain anthrax must be referred to a CLRN laboratory for further characterization in order to comply with the Human Pathogens and Toxin Act (http:// www.phac-aspc.gc.ca/lab-bio/regul/hpta-lapht-eng.php). Similar protocols exist in other countries. Tests for the presence of B. anthracis may be requested for diverse specimens, such as animal products (e.g., wool, hides, hair, and bone meal) from regions of endemicity, soil or other materials from old burial sites or tannery or laboratory sites due for redevelopment, or other environmental materials associated with outbreaks (e.g., sewage sludge). Detection in such specimens may mean searching for rather few spores of B. anthracis among those of many other species, especially other members of the B. cereus group. Some environmental specimens may contain substances that inhibit germination and growth of B. anthracis (69). At present, there is no enrichment method specific for B. anthracis, and culture by the selective agar techniques described above is the best approach.

IDENTIFICATION Species level identification of an aerobic spore-forming organism usually requires examination of microscopic features, colonial morphology, biochemical tests, and increasingly, analysis of nucleic acid or protein profiles. As a first step, it is important to establish that the isolate really is an aerobic endospore-forming organism. Bacillus species are typically described as Gram-positive rods, but some are Gram variable, and Gram positivity is readily lost in older cultures. Some species or strains simply appear Gram negative. Paenibacillus and Brevibacillus are more likely to stain Gram variable or Gram negative. Phase-contrast microscopy and spore staining can be used to distinguish PHB granules and other storage inclusions from true spores. For some species, the storage granules can actually assist with identification. When grown on carbohydrate-rich media, such as glucose nutrient agar, the large cells of B. megaterium may accumu-

451

late PHB (Fig. 2d) and appear vacuolated or foamy. However, the microscopic morphology of the sporangia are much more helpful for distinguishing between species (Fig. 2). Spore shapes vary from cylindrical through ellipsoidal to spherical. Occasionally, bean- or kidney-shaped, curvedcylindrical, and pear-shaped spores are also seen. Spores may be positioned terminally, subterminally, or centrally; the sporangia may appear swollen or distended. Despite within-species and within-strain variation, sporangial morphologies can be sufficiently characteristic to allow an experienced worker to make a tentative identification. For example, B. laterosporus produces very distinctive ellipsoidal spores that have thickened rims on one side, such that they appear to be laterally displaced in the sporangia (Fig. 2j). Recognition of B. thuringiensis is largely dependent on observation of its cuboid or diamond-shaped parasporal crystals in sporulated cultures (Fig. 2c). In general, morphological features of the vegetative cell tend to be less informative, but they can still assist in species identification. Vegetative cells vary in width from about 0.5 to 1.5 μm and in length from 1.5 to 8 μm. They may occur singly or in chains. Usually, cells are round-ended, but some are square or, occasionally, tapered (e.g., P. alvei) (Fig. 2l). Many species are motile. Growth characteristics are often diagnostic. B. subtilis and B. licheniformis exhibit similar colonial and microscopic morphologies but different growth characteristics, being strictly aerobic and facultatively anaerobic, respectively. B. megaterium and B. cereus are both large-celled species, but the former is strictly aerobic while the latter is facultatively anaerobic. Species of aerobic endospore-forming bacteria show a very wide range of colonial morphologies, both within and between species (Fig. 3). After 24 to 48 h of growth, sizes range from 1 to 5 mm. Color commonly ranges from buff or creamy gray to off-white, but some strains may produce orange pigment. Morphology can vary from moist and glossy, through granular, to wrinkled. Shapes can vary from round to irregular, sometimes spreading, with entire, through undulate or crenate, to fimbriate edges. Elevations range from effuse, through raised, to convex. Consistency is usually butyrous, but mucoid and dry, adherent colonies are not uncommon. On blood agar, hemolysis may be absent, slight, marked, partial, or complete. Despite this diversity, Bacillus colonies are not generally difficult to recognize. B. subtilis and B. licheniformis produce similar colonies which are exceptionally variable and often appear to be mixed cultures (Fig. 3j). These colonies are of moderate (2 to 4 mm) diameter, irregular in shape, and range from moist and butyrous or mucoid, with margins varying from undulate to fimbriate through membranous with an underlying mucoid matrix, with or without mucoid beading at the surface, to rough and crusty as they dry. The “licheniform” colonies of B. licheniformis tend to be quite adherent. B. circulans is another heterogeneous species, and in about 13% of strains, rotating and migrating microcolonies, which may show spreading growth, have been observed (Fig. 3k). However, closer examination of the spreading strains has revealed some of them to be genotypically distinct, such that they are now classified as species of Paenibacillus. Despite their variability, colonies of B. cereus are readily recognized. They are characteristically large (2 to 7 mm in diameter) and vary in shape from circular to irregular, with entire to undulate, crenate, or fimbriate edges (Fig. 3b). They usually have matte or granular textures, but smooth and moist colonies are not uncommon. When grown on egg yolk agar, strains of the B. cereus group, including albeit

452 n

BACTERIOLOGY

FIGURE 3 Colonies of endospore-forming bacteria on blood agar (a to i) and nutrient agar (j to l) after 24 to 36 h at 37°C. Bars, 2 mm. (a) B. anthracis; (b) B. cereus; (c) B. thuringiensis; (d) B. megaterium; (e) B. pumilus; (f) Lysinibacillus sphaericus; (g) Brevibacillus brevis; (h) Brevibacillus laterosporus; (i) Paenibacillus polymyxa; (j) B. subtilis; (k) B. circulans; (l) Paenibacillus alvei. doi:10.1128/9781555817381.ch26.f3

to a lesser extent, B. anthracis, synthesize lecithinases, which produce opaque zones of precipitation around colonies (i.e., usually after overnight or perhaps 24 h of incubation). The optimum growth temperature is about 37°C, with minima and maxima of 15 to 20°C and 40 to 45°C, respectively. However, routine, reliable differentiation of individual species within the B. cereus group can be challenging. Features considered characteristics of individual species, including the anthrax toxin and capsule of B. anthracis, and the parasporal crystals of B. thuringiensis, are encoded by plasmids.

The acquisition or loss of a plasmid may alter the features of a particular strain and can undermine identification. Colonies of B. anthracis and B. cereus can be similar in appearance, although those of the former are generally smaller and nonhemolytic and may show more spiking or tailing along the lines of inoculation streaks. Compared with the usually butyrous consistency of B. cereus or B. thuringiensis, colonies of B. anthracis are very tenacious and may be pulled into standing peaks with a loop. B. anthracis colonies are also susceptible to diagnostic gamma phage.

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

B. mycoides produces characteristic rhizoid or hairy-looking, adherent colonies which readily cover the whole agar surface. B. anthracis capsule production is stimulated by plating on nutrient agar containing 0.7% sodium bicarbonate followed by overnight incubation under 5 to 7% CO2 (candle jars perform well). Colonies of the capsulated organism appear mucoid, and the capsule can be visualized by M’Fadyean or India ink staining of smears. Phenotypic tests are widely used for species-level identification. For diagnostic purposes, the aerobic endospore formers comprise two groups: the reactive ones, which give positive results in various routine biochemical tests and which are therefore easier to identify (Table 1), and the nonreactive ones, which give few, if any, positive results in such tests. Nonreactive isolates tend to dominate the identification requests sent to reference laboratories. In addition to the traditional phenotypic testing methods, convenient miniaturized formats such as API (bioMérieux SA, Marcy-l’Etoile, France) and automated identification systems such as VITEK (bioMérieux, Inc., Durham, NC) and Biolog (Hayward, CA) also exist. With such systems, it is

TABLE 1

453

important to keep in mind that, if a species is not included in the database, accurate identification is not possible. These systems also have limited value in distinguishing between some members of the B. cereus group (127), with the exception of the presumptive identification of B. anthracis (128). Both the API 50 CH and API 20 E strips together or the API 50 CH alone, along with API 50 CHB/E medium, can be used for the identification of 38 taxa of Bacillus species and related genera and requires up to 48 h of incubation. A large number of Bacillus and related genera are listed in the Biolog database using the GEN III (63 taxa) or GP2 (53 taxa) microplates. Set-up is minimal, and bacteria can be identified in as little as 2 h. BioMérieux has designed the BCL card for the automated identification of aerobic endospore-forming bacteria of the Bacillaceae family using the VITEK 2 system. A total of 42 species can be identified, and final results are obtained after 14 h (129). The BD Phoenix automated microbiology system (BD Diagnostics, Sparks, MD) lists 12 Bacillus species and related taxa in its Gram-positive panel identification database. The commercially available databases are expanding, and continue to

Characters for differentiating some species of Bacillus, Geobacillus, and Paenibacillusa Bacillus Geobacillus

E E S, T S (C) − + − − + +

E E S, C S (T) + + − − + +

E S, T + − −

v − − w + − − + + − −

− − − − + − − − + − −

+ − − − + v − v v − v

+ + − +/w + − − + v − −

+ + − − +/w − − v v − −

− − − + + − − + + + +

− − − + + − + + − − +

v − − − + − − v v + −

+ − − − + − − − v − v

− v v − + − v

− − − − +/w +/w +

− + − − − v +

− w + v − − +

− − − − w v +

− + + + + + +

− + v − − v v

+ + + + + + +

− + v v + − +

− − + + + v − + (+) − v

+ − +/w

+/w

v − − + + − − + v − −

+ − − w + − − v v − −

− − + − − − +

− + + − − (+) +

− + + − − (+) +

− − − − − + −

− + + + + + +

− + + + + + +

+ − − + + (+) − + + − +

v − − + − − − + − − +

− − + + + v [(−)] − + (+) [+] − +

− + + (+) + + +

− + + − + + +

− + + v + + +

− + − − + + +

− + [v] + [−] − − + [−] +

− + + +

0.8 (+) +

0.8 (+) +

0.8 v +

0.9 − +

P. alvei

− − + + + + − + + − +

v − − + + − − + + − +

B. lentus

− − + + + − − + + − +

v − − + + − − + + − +

B. firmus

E, S E E (C) S, C S, T S, C − + v − − − − + +

B. circulans

E S, C − − +/w

E (C) [E] E S, C S − − − − + +

B. megaterium

E S (C) − − +

C, E S, C − − −

B. cytotoxicus

E (C) S − + +

E E E (C) S, C S, C S, C − − − − − − − − +

0.8 − +

0.8 (−) +

0.7 − +

P. validus

E S, T v − −/w

≥1.0 1.4 (+) + − +

P. macerans

G. thermodenitrificans

E E (S) S, C S, C, T v + − − − +

1.2 + −

P. polymyxa

G. stearothermophilus

0.8 v +

1.2 1.2 + + − +

1.2 + +

B. coagulans

0.9 v +

B. mycoides

0.7 − +

Paenibacillus

B. circulans group

B. thuringiensis

B. pumilus

0.8 (+) +

B. anthracis

B. licheniformis

0.8 (+) +

B. cereusc

B. amyloliquefaciens

0.8 (−) +

Characterb

Rod mean diam (μm) Chains of cells Motility Sporangiad Spore shape Spore position Sporangium swollen Parasporal crystals Anaerobic growth Growth at: 50°C 65°C Egg yolk reaction Casein hydrolysis Starch hydrolysis Arginine dihydrolase Indole production Gelatin hydrolysis Nitrate reduction Gas from carbohydrates Voges-Proskauer Acid from: D-Arabinose Glycerol Glycogen Inulin Mannitol Salicin D-Trehalose

B. cereus group

B. subtilis

B. subtilis group

0.8 − +

a Symbols and abbreviations: +, ≥85% positive; +/w, positive or weakly positive; w, weakly positive; (+), 75 to 84% positive; v, variable (26 to 74% positive); (−), 16 to 25% positive; −, 0 to 15% positive; −/w, negative or weakly positive. b Arginine dihydrolase, indole production, gelatin hydrolysis, and nitrate reduction reactions were determined using tests in the API 20E strip (bioMérieux). Acid from carbohydrate reactions was determined using the API 50CHB System (bioMérieux). c Results shown in brackets are for the biotype isolated particularly in connection with outbreaks of emetic-type food poisoning and for strains of serovars 1, 3, 5, and 8, which are commonly associated with such outbreaks. d Spore shape: C, cylindrical; E, ellipsoidal; S, spherical. Spore position: C, central or paracentral; S, subterminal; T, terminal. The most common shapes and positions are listed first, and those shown in parentheses are infrequently observed.

454 n

BACTERIOLOGY

improve, but keeping up with the changing taxonomy and deluge of new aerobic, endospore-forming species is a significant challenge. To be effective in identifying a particular species, a diagnostic database must account for intraspecies variability. Creation of a robust profile requires multiple authentic strains from a range of sources, yet many new species are proposed on the basis of single isolates. As such, commercial kits should always be used in conjunction with other methods. Nucleic acid-based methods, especially 16S rRNA gene sequencing, may effectively identify strains that cannot be definitively characterized by morphological or biochemical methods. Guidelines for sequencing-based identification of bacteria are outlined in Clinical Laboratory Standards Institute (CLSI) document MM-18A (130). Some species, notably B. cereus, B. thuringiensis, and B. anthracis, are not effectively differentiated by 16S rRNA gene sequencing. B. anthracis-specific single-nucleotide polymorphisms have been identified using the hypervariable 16S-23S internal transcribed spacer region (131), but sequencing of this target for the definitive identification of B. anthracis is not recommended. In the past, there were concerns about the accuracy and availability of reference sequences. However, these have largely been resolved and, for type and reference strains of most species, high quality 16S rRNA gene sequences can be found in public databases (e.g. GenBank). In recent years, there has been growing interest in the use of MALDI-TOF (MS) platforms for bacterial identification. Three commercial systems, the VITEK-MS (bioMérieux), the MALDI Biotyper (Bruker Corp., Billerica, MA), and the Andromas (Andromas SAS, Paris, France), are currently available. Systems are rapid, easy to use, and facilitate identification of isolates cultured on solid media. Initial instrument cost is high; however, reagent costs are minimal. Methods compatible with liquid media (e.g., blood culture bottles) have been described. As with the phenotypic methods, the key limitation to accurate MS-based identification is the availability of robust databases for interpretation of the MALDI-TOF profiles. Results at the genus level only or misidentifications are still common for this group of organisms (132). These databases may not include recent or uncommon species, and profiles for closely related species (e.g., B. subtilis group or B. cereus sensu lato) may be too similar to differentiate (133). However, analytical methods continue to improve, and effective differentiation of B. subtilis from B. amyloliquefaciens and B. cereus (sensu stricto) from B. thuringiensis has been reported (134). Supplementing commercial databases with locally produced profiles using in-house strains may help improve identification (125). Other approaches to identification include chemotaxonomic fingerprinting by fatty acid methyl ester profiling, pyrolysis MS, and Fourier transform infrared spectroscopy (135). All of these approaches have been successfully applied either across genera or to small groups, but as with other profiling methods, large databases of authentic strains are necessary for accurate identification. Some databases, such as the Microbial Identification System software (Microbial ID, Inc., Newark, DE) for fatty acid methyl ester analysis, are commercially available.

variations, but serotyping is not commonly used today. Similarly, protein-based multilocus enzyme electrophoresis has been replaced by nucleic acid-based genotyping methods. Many PCR- and DNA sequencing-based typing methods have been described, but only a few have gained widespread acceptance, including multiple-locus variable-number tandem repeat analysis (MLVA), multilocus sequence typing (MLST), and increasingly, whole-genome sequencing. MLVA is primarily used for typing of B. anthracis. It is a PCR-based fragment analysis method that targets copy number variations among DNA elements located on the B. anthracis chromosome and virulence plasmids. The differences can be resolved by agarose electrophoresis, capillary electrophoresis, or MS (136–139). The MLVA-8 scheme targets eight loci: six chromosomal plus one on each of pXO1 and pXO2 (139). It was the first typing method that could reliably differentiate B. anthracis strains, and during the 2001 biothreat event, MLVA-8 implicated the Ames strains as the cause of the outbreak (114). The resolution of MLVA increases with the number of targets, and schemes that use 15, 25, and 31 loci have been developed (81, 137). MLVAbank (mlva.u-psud.fr/mlvav4/genotyping/index.php) is an online repository of MLVA profiles and a valuable resource for strain comparison. MLST is a DNA sequencing-based method that exploits nucleotide polymorphisms found in sets of “housekeeping” genes. Because DNA sequences are unequivocal and portable, many view MLST as the gold standard method for B. cereus genotyping. Several MLST schemes have been described, and each targets a different set of six to seven genes. Online databases are available for several of these, including the Priest (pubmlst.org/bcereus) and Tourasse-Helgason (mlstoslo.uio.no) schemes (140, 141), the latter of which provides data on any of five published MLTS schemes for B. cereus. The HyperCat database (also at mlstoslo.uio.no) includes typing data for >2,200 strains and allows comparison of MLST, multilocus enzyme electrophoresis, and amplified fragment length polymorphism data (142). Despite some differences among gene targets, the MLST methods provide a consistent view of the B. cereus group. From the genotypic perspective, B. cereus sensu lato is heterogeneous, and correlation between the phylogenetic clusters and traditional, phenotypic divisions is poor. Some analyses indicate that B. pseudomycoides, B. cytotoxicus (143), and B. toyonensis do comprise distinct clades, B. anthracis is recognized as a pathogenic clone, but B. cereus and B. thuringiensis strains are largely indistinguishable. Although reports have suggested that the B. cereus strains associated with emetic food poisoning exhibit distinct characteristics, the emetic toxin can actually be produced by multiple discrete phylogenetic lineages (144– 146). Whole-genome sequencing has become an increasingly affordable and popular tool for strain typing and molecular epidemiology. Whereas MLST may involve sequencing a few thousand nucleotides, the average genomic dataset includes millions of nucleotides. This provides extraordinary resolution and can reveal single nucleotide polymorphisms among strains that, by other typing methods, appear to be identical (147). This approach has been used to examine the population structure of B. cereus sensu lato (148) and the evolution of B. anthracis (149–151). Increasingly, genomic approaches are also being used to examine phylogenetic relationships among other aerobic endospore-forming bacteria (152).

TYPING Typing schemes have been developed only for a few species of aerobic endospore-forming bacteria, most notably the B. cereus group (including B. anthracis). In the past, this group was differentiated into serovars based on flagellar antigen

SEROLOGIC TESTS Serologic tests for anthrax have been developed, but such assays are not available for infections due to other aerobic

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

endospore-forming bacteria. In outbreak situations, serologic tests for anthrax are of limited utility because seroconversion takes time, whereas effective treatment and public health response require rapid diagnostics. Moreover, in human anthrax, early treatment sometimes prevents development of a detectable rise in antibody titer (115). However, serologic assays are valuable for retrospective surveillance and epidemiological investigations and can also be used to monitor vaccination effectiveness. After the 2001 bioterror event, serologic assays aided in the effort to confirm cases (153). The Ascoli test, which dates from 1911, is a precipitin test that uses hyperimmune serum raised to B. anthracis whole-cell antigen to provide rapid, retrospective evidence of infection. Despite its age, the Ascoli test is still used by veterinarians in Eastern Europe and central Asia (154). Anthraxin, a heat-stable extract from a noncapsulated strain of B. anthracis, is the basis for a skin test that has been licensed for human and animal use since 1962. It is widely acclaimed and remains in use in countries of the former USSR (155). If a delayed-type hypersensitivity reaction develops after injection of anthraxin, it is interpreted as indicating cell-mediated immunity to B. anthracis. The test can be used to monitor vaccine-induced immune status after periods of several years or to diagnose anthrax retrospectively. Anthraxin does not contain highly specific antigens and relies on the fact that the only Bacillus species likely to proliferate within and throughout an animal is B. anthracis. The three protein components of the anthrax toxin (PA, LF, and EF), and antibodies to them, can be used in enzyme immunoassay systems. For routine confirmation of anthrax infection or for monitoring response to anthrax vaccines, antibodies against PA alone appear to be satisfactory and have proved useful for epidemiological investigations with humans and animals. PA, LF, and EF are available commercially (List Biological Laboratories, Inc., Campbell, CA; http://www.listlabs.com). In cases where culture has failed, serologic assays for the detection of antibody response against PA have been used, in combination with NAAT or immunohistochemistry test results, to confirm the diagnosis of anthrax. During the 2001 outbreak, a quantitative human anti-PA immunoglobulin G (IgG) enzyme-linked immunosorbent assay, performed at the CDC, was positive only with sera from individuals with anthrax or vaccinated with AVA vaccine (153). A qualitative kit (QuickELISA Anthrax-PA kit) for the detection of anti-PA IgG and IgM antibodies in human serum has been developed by Immunetics (Boston, MA). It is FDA cleared and CE marked in Europe.

ANTIMICROBIAL SUSCEPTIBILITIES The Clinical Laboratory Standards Institute (CLSI) has published approved guidelines describing susceptibility testing methods and suggested agents for primary testing in the case of B. anthracis, as well as interpretive criteria for other Bacillus spp. (156). Comparable information is also included in chapter 74 of this Manual. The recommended procedure is broth microdilution. Agar dilution and Etest may also be performed (157–160). LRN sentinel level laboratories do not and should not perform antimicrobial susceptibility testing of B. anthracis. If B. anthracis cannot be ruled out, the sentinel level laboratory submits the isolate to its designated LRN reference laboratory for confirmatory identification; the latter laboratory will forward the isolate to the federal level (CDC) for antimicrobial susceptibility testing, se-

455

quencing, and archiving. The same is true for sentinel level food and veterinary laboratories.

Bacillus anthracis Most strains of B. anthracis are susceptible to penicillin, although exceptions have been reported (157–159). However, the presence of inducible β-lactamases in some isolates precludes the use of penicillin as a single agent in the treatment of systemic anthrax (161). Strains are also susceptible to ciprofloxacin, clindamycin, chloramphenicol, doxycycline, levofloxacin, gentamicin, tetracycline, tobramycin, and vancomycin (157–160, 162, 163). In contrast, the majority of B. anthracis isolates exhibit reduced susceptibility (or “nonsusceptibility”) to some extended- and broad-spectrum cephalosporins, specifically cefuroxime (157), ceftriaxone (158), and cefotaxime (159), rendering these a poor choice for treatment (158), and in vitro results, even if susceptible, may not predict clinical efficacy. Resistance or reduced susceptibility has also been shown for trimethoprimsulfamethoxazole (TMP-SMX), erythromycin, and azithromycin (158–160). In general, susceptibility testing of B. anthracis is often not required, nor are interpretive breakpoints established (other than susceptible) by the CLSI, with the exception of penicillin (156, 164). Agents for primary testing may include penicillin, doxycycline, tetracycline, and ciprofloxacin and must be performed in a reference laboratory with BSL-3 capacity. Postexposure prophylaxis can prevent inhalational anthrax following exposure to aerosols containing B. anthracis spores. The recommended regimen is 60 days of oral antimicrobial therapy and three doses of AVA (165). When selecting antimicrobial agents, consideration for the possibility of antimicrobial resistance should be given. For adult and pediatric patients, the recommended first-line antimicrobial agent is ciprofloxacin or doxycycline (166). Levofloxacin is recommended as a second-line antimicrobial for adults (165), although there is limited safety data regarding use beyond 28 days. For pregnant women, ciprofloxacin is recommended over doxycycline, though doxycycline may be used if ciprofloxacin is unavailable (167). Amoxicillin may be used if the isolate at issue is susceptible to penicillin. Amoxicillin is also recommended in the treatment of children and lactating women and when other antimicrobial agents are not considered safe. Clindamycin, chloramphenicol, rifampin, vancomycin, and other fluoroquinolones are also suitable for the treatment of patients unable to tolerate recommended antibiotics (165). For severe cases of anthrax (e.g., fulminant bacteremia, inhalational, gastrointestinal, or injectional anthrax), there is often meningeal involvement and the recommended treatment is intravenous ciprofloxacin for 7 to 10 days plus one or two additional drugs (65, 165). In the presence of inflammation, central nervous system penetration of ciprofloxacin is much higher than that of doxycycline. For cases of uncomplicated, naturally acquired, cutaneous anthrax, a 7- to 10-day course of oral ciprofloxacin or doxycycline is recommended, but penicillin V or amoxicillin may also be used if susceptibility is confirmed. Bioterrorismrelated cutaneous anthrax should be subject to treatment as described for postexposure prophylaxis due to the risk of aerosol exposure.

Bacillus cereus Group (Not B. anthracis) There have been rather few studies of the antimicrobial susceptibility of B. cereus, and most information has to be gleaned from reports of individual cases or outbreaks. B. cereus and B. thuringiensis produce penicillinases and a

456 n

BACTERIOLOGY

broad-spectrum β-lactamase and are thus resistant to penicillins, cephalosporins, and β-lactamase inhibitor combinations (160, 168). Moderate or intermediate susceptibility has been demonstrated for TMP-SMX, clindamycin, and tetracycline in a portion of isolates (160, 168). All strains are susceptible to imipenem and vancomycin, and most are susceptible to chloramphenicol, erythromycin, gentamicin, ciprofloxacin, and daptomycin (160, 168, 169). Despite this, treatment failures have been reported, even for regimens containing vancomycin. Such cases were summarized in the previous version of this chapter (170). Oral ciprofloxacin has been used successfully in the treatment of B. cereus wound infections, bacteremia, and pulmonary infection. Clindamycin with gentamicin, given early, appears to be the best treatment for ophthalmic infections caused by B. cereus (171). Recommended agents for primary susceptibility testing of Bacillus spp. (not B. anthracis) include vancomycin, fluoroquinolones, and clindamycin, and interpretive criteria are available for many antibiotics (156).

foodstuffs by aerobic endospore-forming bacteria is common, as is transient and asymptomatic fecal carriage. Ideally, to establish that a putative pathogen is the etiological agent of a food poisoning event, the organism should be isolated from the epidemiologically incriminated food in significant numbers (>105 CFU/g) and the same strain (genotype, biovar, or plasmid type, etc.) should also be isolated in significant numbers from acute-phase specimens (feces or vomitus) obtained from patients but not from healthy controls. For events associated with B. cereus, it should also be possible to detect emetic toxin and/or enterotoxin in the food. In cases where clinical significance is suspected or established, but the isolate does not represent a “usual suspect,” it is preferable that the identification be confirmed by DNA sequencing. Ideally, nucleotide sequences associated with rare and potentially novel species should be submitted to public databases, where they will be accessible to other laboratories searching for information about the same aerobic endospore-forming organism.

Other Species REFERENCES There is a paucity of antimicrobial susceptibility data for other Bacillus species and related genera, due to the low frequency at which these are found to be clinically significant. Most of the available information has been derived from individual clinical case studies, where antibiotics were administered empirically and treatment may or may not have been guided by in vitro susceptibility results. As environmental organisms, Bacillus spp. may exhibit a wide array of resistance and susceptibility patterns against commonly used antibiotics (169). When isolated from normally sterile sites and clinically warranted, the laboratory may use the published CLSI approved guidelines (156). The antimicrobial agents suggested for primary testing are vancomycin, clindamycin, and fluoroquinolones. When tested, clinical isolates of non-B. cereus, non-B. anthracis species are typically susceptible to vancomycin, daptomycin, gentamicin, ciprofloxacin, imipenem, erythromycin, and TMP-SMX (168). Variable susceptibility exists for penicillins, cephalosporins, chloramphenicol, clindamycin, and tetracycline (168). Vancomycin resistance is rare, but it has been reported (172, 173).

EVALUATION, INTERPRETATION, AND REPORTING OF RESULTS The isolation of B. anthracis is always significant and requires urgent reporting, but the majority of aerobic endosporeforming bacteria are innocuous environmental organisms, not professional pathogens. Although frequently encountered in the microbiology laboratory, most isolates are of no clinical relevance. However, opportunistic infections have been reported since the late 19th century, and over the past 30 years, the clinical importance of aerobic endospore formers (most, but not all, of them Bacillus species) has become widely accepted. One must be wary of dismissing any organism as a mere contaminant before considering the clinical context of its isolation. If it is obtained in pure culture (or at least appears to dominate the microbiota), if it is isolated in large numbers, or if it is isolated from more than one clinical specimen, then the organism should be considered of potential clinical significance. Moderate or heavy growth of bacilli from wounds is usually significant, and B. cereus infections of the eye are serious emergencies that should always be reported immediately to the physician. In food-borne illness investigations, qualitative isolation tests are insufficient because low-level contamination of

1. Logan NA, Berge O, Bishop AH, Busse HJ, De Vos P, Fritze D, Heyndrickx M, Kampfer P, Rabinovitch L, Salkinoja-Salonen MS, Seldin L, Ventosa A. 2009. Proposed minimal standards for describing new taxa of aerobic, endospore-forming bacteria. Int J Syst Evol Microbiol 59:2114–2121. 2. Bhandari V, Ahmod NZ, Shah HN, Gupta RS. 2013. Molecular signatures for Bacillus species: demarcation of the Bacillus subtilis and Bacillus cereus clades in molecular terms and proposal to limit the placement of new species into the genus Bacillus. Int J Syst Evol Microbiol 63:2712– 2726. 3. Castagnola E, Fioredda F, Barretta MA, Pescetto L, Garaventa A, Lanino E, Micalizzi C, Giacchino R, Dini G. 2001. Bacillus sphaericus bacteraemia in children with cancer: case reports and literature review. J Hosp Infect 48:142– 145. 4. Berry C. 2012. The bacterium, Lysinibacillus sphaericus, as an insect pathogen. J Invertebr Pathol 109:1–10. 5. Ash C, Priest FG, Collins MD. 1993. Molecular identification of rRNA group 3 bacilli (Ash, Farrow, Wallbanks and Collins) using a PCR probe test: proposal for the creation of a new genus Paenibacillus. Antonie van Leewenhoek 64:253– 260. 6. Ko KS, Kim YS, Lee MY, Shin SY, Jung DS, Peck KR, Song JH. 2008. Paenibacillus konsidensis sp. nov., isolated from a patient. Int J Syst Evol Microbiol 58:2164–2168. 7. Roux V, Fenner L, Raoult D. 2008. Paenibacillus provencensis sp. nov., isolated from human cerebrospinal fluid, and Paenibacillus urinalis sp. nov., isolated from human urine. Int J Syst Evol Microbiol 58:682–687. 8. Roux V, Raoult D. 2004. Paenibacillus massiliensis sp. nov., Paenibacillus sanguinis sp. nov. and Paenibacillus timonensis sp. nov., isolated from blood cultures. Int J Syst Evol Microbiol 54:1049–1054. 9. Ash C, Farrow JAE, Wallbanks S, Collins MD. 1991. Phylogenetic heterogeneity of the genus Bacillus revealed by comparative analysis of small-subunit-ribosomal RNA sequences. Lett Appl Microbiol 13:202–206. 10. Logan NA, Forsyth G, Lebbe L, Goris J, Heyndrickx M, Balcaen A, Verhelst A, Falsen E, Ljungh A, Hansson HB, De Vos P. 2002. Polyphasic identification of Bacillus and Brevibacillus strains from clinical, dairy and industrial specimens and proposal of Brevibacillus invocatus sp. nov. Int J Syst Evol Microbiol 52:953–966. 11. Parvez N, Cornelius LK, Fader R. 2009. Brevibacillus brevis peritonitis. Am J Med Sci 337:297–299.

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n 12. de Oliveira EJ, Rabinovitch L, Monnerat RG, Passos LK, Zahner V. 2004. Molecular characterization of Brevibacillus laterosporus and its potential use in biological control. Appl Environ Microbiol 70:6657–6664. 13. Yabbara KF, Juffali F, Matossian RM. 1977. Bacillus laterosporus endophthalmitis. Arch Ophthalmol 95:2187–2189. 14. Saile E, Koehler TM. 2006. Bacillus anthracis multiplication, persistence, and genetic exchange in the rhizosphere of grass plants. Appl Environ Microbiol 72:3168–3174. 15. Dey R, Hoffman PS, Glomski IJ. 2012. Germination and amplification of anthrax spores by soil-dwelling amoebas. Appl Environ Microbiol 78:8075–8081. 16. Jeon YL, Yang JJ, Kim MJ, Lim G, Cho SY, Park TS, Suh JT, Park YH, Lee MS, Kim SC, Lee HJ. 2012. Combined Bacillus licheniformis and Bacillus subtilis infection in a patient with oesophageal perforation. J Med Microbiol 61:1766–1769. 17. Logan NA. 1988. Bacillus species of medical and veterinary importance. J Med Microbiol 25:157–165. 18. Beebe JL, Koneman EW. 1995. Recovery of uncommon bacteria from blood: association with neoplastic disease. Clin Microbiol Rev 8:336–356. 19. Thomas M, Whittet H. 1991. Atypical meningitis complicating a penetrating head injury. J Neurol Neurosurg Psychiatry 54:92–93. 20. Logan NA. 2012. Bacillus and relatives in foodborne illness. J Appl Microbiol 112:417–429. 21. Stickel F, Droz S, Patsenker E, Bogli-Stuber K, Aebi B, Leib SL. 2009. Severe hepatotoxicity following ingestion of Herbalife nutritional supplements contaminated with Bacillus subtilis. J Hepatol 50:111–117. 22. Spinosa MR, Wallet F, Courcol RJ, Oggioni MR. 2000. The trouble in tracing opportunistic pathogens: cholangitis due to Bacillus in a French hospital caused by a strain related to an Italian probiotic? Microb Ecol Health Dis 12:99–101. 23. Idelevich EA, Pogoda CA, Ballhausen B, Wullenweber J, Eckardt L, Baumgartner H, Waltenberger J, Peters G, Becker K. 2013. Pacemaker lead infection and related bacteraemia caused by normal and small colony variant phenotypes of Bacillus licheniformis. J Med Microbiol 62:940– 944. 24. Ozkocaman V, Ozcelik T, Ali R, Ozkalemkas F, Ozkan A, Ozakin C, Akalin H, Ursavas A, Coskun F, Ener B, Tunali A. 2006. Bacillus spp. among hospitalized patients with haematological malignancies: clinical features, epidemics and outcomes. J Hosp Infect 64:169–176. 25. Haydushka IA, Markova N, Kirina V, Atanassova M. 2012. Recurrent sepsis due to Bacillus licheniformis. J Glob Infect Dis 4:82–83. 26. Hannah WN Jr, Ender PT. 1999. Persistent Bacillus licheniformis bacteremia associated with an international injection of organic drain cleaner. Clin Infect Dis 29:659–661. 27. Galanos J, Perera S, Smith H, O’Neal D, Sheorey H, Waters M. 2003. Bacteremia due to three Bacillus species in a case of Munchausen’s syndrome. J Clin Microbiol 41:2247–2248. 28. Mikkola R, Kolari M, Andersson MA, Helin J, SalkinojaSalonen MS. 2000. Toxic lactonic lipopeptide from food poisoning isolates of Bacillus licheniformis. Eur J Biochem 267:4068–4074. 29. Kimouli M, Vrioni G, Papadopoulou M, Koumaki V, Petropoulou D, Gounaris A, Friedrich AW, Tsakris A. 2012. Two cases of severe sepsis caused by Bacillus pumilus in neonatal infants. J Med Microbiol 61:596–599. 30. Farhat H, Chachaty E, Antoun S, Nitenberg G, Zahar JR. 2008. Two cases of Bacillus infection and immunodepression. Med Mal Infect 38:612–614. 31. From C, Hormazabal V, Granum PE. 2007. Food poisoning associated with pumilacidin-producing Bacillus pumilus in rice. Int J Food Microbiol 115:319–324.

457

32. Beesley CA, Vanner CL, Helsel LO, Gee JE, Hoffmaster AR. 2010. Identification and characterization of clinical Bacillus spp. isolates phenotypically similar to Bacillus anthracis. FEMS Microbiol Lett 313:47–53. 33. Duncan KO, Smith TL. 2011. Primary cutaneous infection with Bacillus megaterium mimicking cutaneous anthrax. J Am Acad Dermatol 65:e60–e61. 34. Berry N, Hassan I, Majumdar S, Vardhan A, McEwen A, Gokal R. 2004. Bacillus circulans peritonitis in a patient treated with CAPD. Perit Dial Int 24:488–489. 35. Ferrand J, Hadou T, Selton-Suty C, Goehringer F, Sadoul N, Alauzet C, Lozniewski A. 2013. Cardiac device-related endocarditis caused by Paenibacillus glucanolyticus. J Clin Microbiol 51:3439–3442. 36. Kampfer P, Rossello-Mora R, Falsen E, Busse HJ, Tindall BJ. 2006. Cohnella thermotolerans gen. nov., sp. nov., and classification of ‘Paenibacillus hongkongensis’ as Cohnella hongkongensis sp. nov. Int J Syst Evol Microbiol 56:781–786. 37. Ko KS, Oh WS, Lee MY, Lee JH, Lee H, Peck KR, Lee NY, Song JH. 2006. Bacillus infantis sp. nov. and Bacillus idriensis sp. nov., isolated from a patient with neonatal sepsis. Int J Syst Evol Microbiol 56:2541–2544. 38. Ouyang J, Pei Z, Lutwick L, Dalal S, Yang L, Cassai N, Sandhu K, Hanna B, Wieczorek RL, Bluth M, Pincus MR. 2008. Paenibacillus thiaminolyticus: a new cause of human infection, inducing bacteremia in a patient on hemodialysis. Ann Clin Lab Sci 38:393–400. 39. Glazunova OO, Raoult D, Roux V. 2006. Bacillus massiliensis sp. nov., isolated from cerebrospinal fluid. Int J Syst Evol Microbiol 56:1485–1488. 40. Bosshard PP, Zbinden R, Altwegg M. 2002. Paenibacillus turicensis sp. nov., a novel bacterium harbouring heterogeneities between 16S rRNA genes. Int J Syst Evol Microbiol 52:2241–2249. 41. Anikpeh YF, Keller P, Bloemberg GV, Grunenfelder J, Zinkernagel AS. 2010. Spacecraft bacterium, Paenibacillus pasadenensis, causing wound infection in humans. BMJ Case Rep doi:10.1136/bcr.06.2010.3058. 42. Glaeser SP, Falsen E, Busse HJ, Kampfer P. 2013. Paenibacillus vulneris sp. nov., isolated from a necrotic wound. Int J Syst Evol Microbiol 63:777–782. 43. Miller JJ, Scott IU, Flynn HW Jr, Smiddy WE, Murray TG, Berrocal A, Miller D. 2008. Endophthalmitis caused by Bacillus species. Am J Ophthalmol 145:883–888. 44. Szkaradkiewicz A, Chudzicka-Strugala I, Karpinski TM, Goslinska-Pawlowska O, Tulecka T, Chudzicki W, Szkaradkiewicz AK, Zaba R. 2012. Bacillus oleronius and Demodex mite infestation in patients with chronic blepharitis. Clin Microbiol Infect 18:1020–1025. 45. Frankland GC, Frankland PF. 1887. Studies on some new micro-organisms obtained from air. Phil Trans R Soc B 178:257–287. 46. Ceuppens S, Boon N, Uyttendaele M. 2013. Diversity of Bacillus cereus group strains is reflected in their broad range of pathogenicity and diverse ecological lifestyles. FEMS Microbiol Ecol 84:433–450. 47. Drobniewski FA. 1993. Bacillus cereus and related species. Clin Microbiol Rev 6:324–338. 48. Bottone EJ. 2010. Bacillus cereus, a volatile human pathogen. Clin Microbiol Rev 23:382–398. 49. Dolan SA, Littlehorn C, Glode MP, Dowell E, Xavier K, Nyquist AC, Todd JK. 2012. Association of Bacillus cereus infection with contaminated alcohol prep pads. Infect Control Hosp Epidemiol 33:666–671. 50. Balm MN, Jureen R, Teo C, Yeoh AE, Lin RT, Dancer SJ, Fisher DA. 2012. Hot and steamy: outbreak of Bacillus cereus in Singapore associated with construction work and laundry practices. J Hosp Infect 81:224–230. 51. Cluck D, Williamson JC, Glasgo M, Diekema D, Sherertz R. 2012. Bacterial contamination of an automated

458 n

52.

53. 54.

55.

56.

57.

58.

59.

60. 61.

62.

63. 64.

65.

66.

67.

68.

69. 70.

71.

72.

BACTERIOLOGY

pharmacy robot used for intravenous medication preparation. Infect Control Hosp Epidemiol 33:517–520. McLauchlin J, Salmon JE, Ahmed S, Brazier JS, Brett MM, George RC, Hood J. 2002. Amplified fragment length polymorphism (AFLP) analysis of Clostridium novyi, C. perfringens and Bacillus cereus isolated from injecting drug users during 2000. J Med Microbiol 51:990–1000. Durand ML. 2013. Endophthalmitis. Clin Microbiol Infect 19:227–234. Pinna A, Sechi LA, Zanetti S, Usai D, Delogu G, Cappuccinelli P, Carta F. 2001. Bacillus cereus keratitis associated with contact lens wear. Ophthalmology 108:1830–1834. Donzis PB, Mondino BJ, Weissman BA. 1988. Bacillus keratitis associated with contaminated contact lens care systems. Am J Ophthalmol 105:195–197. Bennett SD, Walsh KA, Gould LH. 2013. Foodborne disease outbreaks caused by Bacillus cereus, Clostridium perfringens, and Staphylococcus aureus—United States, 19982008. Clin Infect Dis 57:425–433. Bartoszewicz M, Hansen BM, Swiecicka I. 2008. The members of the Bacillus cereus group are commonly present contaminants of fresh and heat-treated milk. Food Microbiol 25:588–596. Ivy RA, Ranieri ML, Martin NH, den Bakker HC, Xavier BM, Wiedmann M, Boor KJ. 2012. Identification and characterization of psychrotolerant sporeformers associated with fluid milk production and processing. Appl Environ Microbiol 78:1853–1864. Koch R. 1877. Verfahren zur Untersuchung, zum Conservieren und Photogaphiren der Bakterien. Beitr. Biol. Pflanzen 2:399–434. Beyer W, Turnbull PC. 2009. Anthrax in animals. Mol Aspects Med 30:481–489. Fasanella A, Galante D, Garofolo G, Jones MH. 2010. Anthrax undervalued zoonosis. Vet Microbiol 140:318– 331. Vijaikumar M, Thappa DM, Karthikeyan K. 2002. Cutaneous anthrax: an endemic outbreak in south India. J Trop Pediatr 48:225–226. Yakupogullari Y, Koroglu M. 2007. Nosocomial spread of Bacillus anthracis. J Hosp Infect 66:401–402. Meselson M, Guillemin J, Hugh-Jones M, Langmuir A, Popova I, Shelokov A, Yampolskaya O. 1994. The Sverdlovsk anthrax outbreak of 1979. Science 266:1202–1208. Sweeney DA, Hicks CW, Cui X, Li Y, Eichacker PQ. 2011. Anthrax infection. Am J Respir Crit Care Med 184:1333–1341. Bravata DM, Holty JE, Wang E, Lewis R, Wise PH, McDonald KM, Owens DK. 2007. Inhalational, gastrointestinal, and cutaneous anthrax in children. Arch Pediatr Adolesc Med 161:896–905. Meaney-Delman D, Zotti ME, Rasmussen SA, Strasser S, Shadomy S, Turcios-Ruiz RM, Wendel GD Jr, Treadwell TA, Jamieson DJ. 2012. Anthrax cases in pregnant and postpartum women: a systematic review. Obstet Gynecol 120:1439–1449. Centers for Disease Control and Prevention. 2013. Summary of notifiable diseases - United States, 2011. MMWR Morb Mortal Wkly Rep 60:1–120. Turnbull P. 2008. Anthrax in Humans and Animals, 4th ed. World Health Organization, Geneva, Switzerland. Kracalik I, Malania L, Tsertsvadze N, Manvelyan J, Bakanidze L, Imnadze P, Tsanava S, Blackburn JK. 2014. Human cutaneous anthrax, Georgia 2010-2012. Emerg Infect Dis 20:261–264. Bradaric´a N, Punda-Polic´ V. 1992. Cutaneous anthrax due to penicillin-resistant Bacillus anthracis transmitted by an insect bite. Lancet 340:306–307. Fasanella A, Garofolo G, Galella M, Troiano P, De Stefano C, Pace L, Aceti A, Serrecchia L, Adone R. 2013.

73. 74.

75.

76.

77. 78.

79.

80.

81.

82.

83.

84.

85. 86.

87.

88.

Suspect vector transmission of human cutaneous anthrax during an animal outbreak in Southern Italy. Vector Borne Zoonotic Dis 13:769–771. Sirisanthana T, Brown AE. 2002. Anthrax of the gastrointestinal tract. Emerg Infect Dis 8:649–651. Beatty ME, Ashford DA, Griffin PM, Tauxe RV, Sobel J. 2003. Gastrointestinal anthrax. Arch Intern Med 163:2527– 2531. Brachman P, Kaufmann A. 1998. Anthrax, p 95–107. In Evans A, Brachman P (ed), Bacterial Infections of Humans. Plenum Medical Book Company, New York, NY. Jernigan DB, Raghunathan PL, Bell BP, Brechner R, Bresnitz EA, Butler JC, Cetron M, Cohen M, Doyle T, Fischer M, Greene C, Griffith KS, Guarner J, Hadler JL, Hayslett JA, Meyer R, Petersen LR, Phillips M, Pinner R, Popovic T, Quinn CP, Reefhuis J, Reissman D, Rosenstein N, Schuchat A, Shieh WJ, Siegal L, Swerdlow DL, Tenover FC, Traeger M, Ward JW, Weisfuse I, Wiersma S, Yeskey K, Zaki S, Ashford DA, Perkins BA, Ostroff S, Hughes J, Fleming D, Koplan JP, Gerberding JL, National Anthrax Epidemiologic Investigation Team. 2002. Investigation of bioterrorism-related anthrax, United States, 2001: epidemiologic findings. Emerg Infect Dis 8:1019–1028. Frazier AA, Franks TJ, Galvin JR. 2006. Inhalational anthrax. J Thorac Imaging 21:252–258. Ringertz SH, Høiby EA, Jensenius M, Caugant JMDA, Myklebust A, Fossum K. 2000. Injectional anthrax in a heroin skin-popper. Lancet 356:1574–1575. Booth MG, Hood J, Brooks TJ, Hart A, Health Protection Scotland Anthrax Clinical Network. 2010. Anthrax infection in drug users. Lancet 375:1345–1346. Team NAOC. 2011. An Outbreak of Anthrax among Drug Users in Scotland, December 2009 to December 2010. Health Protection Scotland, NHS National Services Scotland, Glasgow, United Kingdom. Grunow R, Klee SR, Beyer W, George M, Grunow D, Barduhn A, Klar S, Jacob D, Elschner M, Sandven P, Kjerulf A, Jensen JS, Cai W, Zimmermann R, Schaade L. 2013. Anthrax among heroin users in Europe possibly caused by same Bacillus anthracis strain since 2000. Euro Surveill 18:20437. Hanczaruk M, Reischl U, Holzmann T, Frangoulidis D, Wagner DM, Keim PS, Antwerpen MH, Meyer H, Grass G. 2014. Injectional anthrax in heroin users, Europe, 20002012. Emerg Infect Dis 20:322–323. Marston CK, Allen CA, Beaudry J, Price EP, Wolken SR, Pearson T, Keim P, Hoffmaster AR. 2011. Molecular epidemiology of anthrax cases associated with recreational use of animal hides and yarn in the United States. PLoS One 6:e28274. Anaraki S, Addiman S, Nixon G, Krahe D, Ghosh R, Brooks T, Lloyd G, Spencer R, Walsh A, McCloskey B, Lightfoot N. 2008. Investigations and control measures following a case of inhalation anthrax in East London in a drum maker and drummer, October 2008. Euro Surveill 13:19076. Eurosurveillance Editorial Team. 2006. Probable human anthrax death in Scotland. Euro Surveill 11:E060817.2. Griffith J, Blaney D, Shadomy S, Lehman M, Pesik N, Tostenson S, Delaney L, Tiller R, DeVries A, Gomez T, Sullivan M, Blackmore C, Stanek D, Lynfield R, Anthrax Investigation Team. 2014. Investigation of inhalation anthrax case, United States. Emerg Infect Dis 20:280–283. Sykes A, Brooks T, Dusmet M, Nicholson AG, Hansell DM, Wilson R. 2013. Inhalational anthrax in a vaccinated soldier. Eur Respir J 42:285–287. Wilson MK, Vergis JM, Alem F, Palmer JR, Keane-Myers AM, Brahmbhatt TN, Ventura CL, O’Brien AD. 2011. Bacillus cereus G9241 makes anthrax toxin and capsule

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

100. 101.

102.

103.

like highly virulent B. anthracis Ames but behaves like attenuated toxigenic nonencapsulated B. anthracis Sterne in rabbits and mice. Infect Immun 79:3012–3019. Klee SR, Brzuszkiewicz EB, Nattermann H, Bruggemann H, Dupke S, Wollherr A, Franz T, Pauli G, Appel B, Liebl W, Couacy-Hymann E, Boesch C, Meyer FD, Leendertz FH, Ellerbrok H, Gottschalk G, Grunow R, Liesegang H. 2010. The genome of a Bacillus isolate causing anthrax in chimpanzees combines chromosomal properties of B. cereus with B. anthracis virulence plasmids. PLoS One 5:e10986. Ehling-Schulz M, Fricker M, Grallert H, Rieck P, Wagner M, Scherer S. 2006. Cereulide synthetase gene cluster from emetic Bacillus cereus: structure and location on a mega virulence plasmid related to Bacillus anthracis toxin plasmid pXO1. BMC Microbiol 6:20. Guinebretiere M-H, Velge P, Couvert O, Carlin F, Debuyser M-L, Nguyen-The C. 2010. Ability of Bacillus cereus group strains to cause food poisoning varies according to phylogenetic affiliation (groups I to VII) rather than species affiliation. J Clin Microbiol 48:3388–3391. Stenfors Arnesen LP, Fagerlund A, Granum PE. 2008. From soil to gut: Bacillus cereus and its food poisoning toxins. FEMS Microbiol Rev 32:579–606. Senesi S, Ghelardi E. 2010. Production, secretion and biological activity of Bacillus cereus enterotoxins. Toxins (Basel) 2:1690–1703. Lapidus A, Goltsman E, Auger S, Galleron N, Segurens B, Dossat C, Land ML, Broussolle V, Brillard J, Guinebretiere MH, Sanchis V, Nguen-The C, Lereclus D, Richardson P, Wincker P, Weissenbach J, Ehrlich SD, Sorokin A. 2008. Extending the Bacillus cereus group genomics to putative food-borne pathogens of different toxicity. Chem Biol Interact 171:236–249. Fagerlund A, Ween O, Lund T, Hardy SP, Granum PE. 2004. Genetic and functional analysis of the cytK family of genes in Bacillus cereus. Microbiology 150:2689– 2697. Ramarao N, Sanchis V. 2013. The pore-forming haemolysins of Bacillus cereus: a review. Toxins (Basel) 5:1119– 1139. Clair G, Roussi S, Armengaud J, Duport C. 2010. Expanding the known repertoire of virulence factors produced by Bacillus cereus through early secretome profiling in three redox conditions. Mol Cell Proteomics 9:1486– 1498. Gohar M, Faegri K, Perchat S, Ravnum S, Okstad OA, Gominet M, Kolsto AB, Lereclus D. 2008. The PlcR virulence regulon of Bacillus cereus. PLoS One 3:e2793. Sanahuja G, Banakar R, Twyman RM, Capell T, Christou P. 2011. Bacillus thuringiensis: a century of research, development and commercial applications. Plant Biotechnol J 9:283–300. Siegel JP. 2001. The mammalian safety of Bacillus thuringiensis-based insecticides. J Invertebr Pathol 77:13–21. Centers for Disease Control and Prevention and National Institutes of Health. 2009. Biosafety in Microbiological and Biomedical Laboratories, 5th ed. US Government Printing Office, Washington, DC. Public Health Agency of Canada and Canadian Food Inspection Agency. 2013. Canadian Biosafety Standards and Guidelines, 1st ed. Public Health Agency of Canada, Ottawa, Ontario, Canada. Baron EJ, Miller JM, Weinstein MP, Richter SS, Gilligan PH, Thomson RB Jr, Bourbeau P, Carroll KC, Kehl SC, Dunne WM, Robinson-Dunn B, Schwartzman JD, Chapin KC, Snyder JW, Forbes BA, Patel R, Rosenblatt JE, Pritt BS. 2013. A guide to utilization of the microbiology laboratory for diagnosis of infectious diseases: 2013 recommendations by the Infectious Diseases Society of

104.

105.

106.

107.

108. 109. 110.

111.

112.

113.

114.

115.

116.

117.

118.

119.

120.

459

America (IDSA) and the American Society for Microbiology (ASM). Clin Infect Dis 57:e22–e121. Centers for Disease Control and Prevention. 2010. Use of anthrax vaccine in the United States. Recommendations of the Advisory Committee on Immunization Practices (ACIP), 2009. MMWR Recommend Rep 59:1–30. Shieh WJ, Guarner J, Paddock C, Greer P, Tatti K, Fischer M, Layton M, Philips M, Bresnitz E, Quinn CP, Popovic T, Perkins BA, Zaki SR, Anthrax Bioterrorism Investigation Team. 2003. The critical role of pathology in the investigation of bioterrorism-related cutaneous anthrax. Am J Pathol 163:1901–1910. Kim J, Yoon MY. 2010. Recent advances in rapid and ultrasensitive biosensors for infectious agents: lesson from Bacillus anthracis diagnostic sensors. Analyst 135:1182– 1190. Pipeline and Hazardous Materials Safety Administration. 2006. Transporting Infectious Substances Safely. Pipeline and Hazardous Materials Safety Administration, US Department of Transportation, Washington, DC. Cybulski RJ Jr, Sanz P, O’Brien AD. 2009. Anthrax vaccination strategies. Mol Aspects Med 30:490–502. Schaeffer AB, Fulton MD. 1933. A simplified method of staining endospores. Science 77:194. Kolsto AB, Tourasse NJ, Okstad OA. 2009. What sets Bacillus anthracis apart from other Bacillus species? Annu Rev Microbiol 63:451–476. Andersson MA, Hakulinen P, Honkalampi-Hamalainen U, Hoornstra D, Lhuguenot JC, Maki-Paakkanen J, Savolainen M, Severin I, Stammati AL, Turco L, Weber A, von Wright A, Zucco F, Salkinoja-Salonen M. 2007. Toxicological profile of cereulide, the Bacillus cereus emetic toxin, in functional assays with human, animal and bacterial cells. Toxicon 49:351–367. Yamaguchi M, Kawai T, Kitagawa M, Kumeda Y. 2013. A new method for rapid and quantitative detection of the Bacillus cereus emetic toxin cereulide in food products by liquid chromatography-tandem mass spectrometry analysis. Food Microbiol 34:29–37. Ehling-Schulz M, Messelhausser U. 2013. Bacillus “next generation” diagnostics: moving from detection toward subtyping and risk-related strain profiling. Front Microbiol 4:32. Hoffmaster AR, Fitzgerald CC, Ribot E, Mayer LW, Popovic T. 2002. Molecular subtyping of Bacillus anthracis and the 2001 bioterrorism-associated anthrax outbreak, United States. Emerg Infect Dis 8:1111–1116. Quinn CP, Dull PM, Semenova V, Li H, Crotty S, Taylor TH, Steward-Clark E, Stamey KL, Schmidt DS, Stinson KW, Freeman AE, Elie CM, Martin SK, Greene C, Aubert RD, Glidewell J, Perkins BA, Ahmed R, Stephens DS. 2004. Immune responses to Bacillus anthracis protective antigen in patients with bioterrorismrelated cutaneous or inhalation anthrax. J Infect Dis 190:1228–1236. Ezzell JW Jr, Abshire TG, Little SF, Lidgerding BC, Brown C. 1990. Identification of Bacillus anthracis by using monoclonal antibody to cell wall galactose-N-acetylglucosamine polysaccharide. J Clin Microbiol 28:223– 231. Muller JD, Wilks CR, O’Riley KJ, Condron RJ, Bull R, Mateczun A. 2004. Specificity of an immunochromatographic test for anthrax. Aust Vet J 82:220–222. Rao SS, Mohan KV, Atreya CD. 2010. Detection technologies for Bacillus anthracis: prospects and challenges. J Microbiol Methods 82:1–10. Irenge LM, Gala JL. 2012. Rapid detection methods for Bacillus anthracis in environmental samples: a review. Appl Microbiol Biotechnol 93:1411–1422. Tang S, Moayeri M, Chen Z, Harma H, Zhao J, Hu H, Purcell RH, Leppla SH, Hewlett IK. 2009. Detection

460 n

121.

122.

123.

124.

125.

126.

127.

128.

129.

130.

131.

132.

133.

134.

135.

BACTERIOLOGY

of anthrax toxin by an ultrasensitive immunoassay using europium nanoparticles. Clin Vaccine Immunol 16:408– 413. Boyer AE, Gallegos-Candela M, Lins RC, Kuklenyik Z, Woolfitt A, Moura H, Kalb S, Quinn CP, Barr JR. 2011. Quantitative mass spectrometry for bacterial protein toxins—a sensitive, specific, high-throughput tool for detection and diagnosis. Molecules 16:2391–2413. Laue M, Fulda G. 2013. Rapid and reliable detection of bacterial endospores in environmental samples by diagnostic electron microscopy combined with X-ray microanalysis. J Microbiol Methods 94:13–21. Dybwad M, van der Laaken AL, Blatny JM, Paauw A. 2013. Rapid identification of Bacillus anthracis spores in suspicious powder samples by using matrix-assisted laser desorption ionization-time of flight mass spectrometry (MALDI-TOF MS). Appl Environ Microbiol 79:5372– 5383. Lasch P, Beyer W, Nattermann H, Stammler M, Siegbrecht E, Grunow R, Naumann D. 2009. Identification of Bacillus anthracis by using matrix-assisted laser desorption ionization-time of flight mass spectrometry and artificial neural networks. Appl Environ Microbiol 75:7229– 7242. Tracz DM, McCorrister SJ, Westmacott GR, Corbett CR. 2013. Effect of gamma radiation on the identification of bacterial pathogens by MALDI-TOF MS. J Microbiol Methods 92:132–134. Schofield DA, Sharp NJ, Vandamm J, Molineux IJ, Spreng KA, Rajanna C, Westwater C, Stewart GC. 2013. Bacillus anthracis diagnostic detection and rapid antibiotic susceptibility determination using ‘bioluminescent’ reporter phage. J Microbiol Methods 95:156–161. Logan NA, Berkeley RCW. 1984. Identification of Bacillus strains using the API system. J Gen Microbiol 130:1871–1882. Logan NA, Carman JA, Melling J, Berkeley RC. 1985. Identification of Bacillus anthracis by API tests. J Med Microbiol 20:75–85. Halket G, Dinsdale AE, Logan NA. 2010. Evaluation of the VITEK2 BCL card for identification of Bacillus species and other aerobic endosporeformers. Lett Appl Microbiol 50:120–126. Clinical and Laboratory Standards Institute. 2008. Interpretive Criteria for Identification of Bacteria and Fungi by DNA Target Sequencing; Approved Guideline. CLSI Document MM18-A. Clinical Laboratory Standards Institute, Wayne, PA. Cherif A, Borin S, Rizzi A, Ouzari H, Boudabous A, Daffonchio D. 2003. Bacillus anthracis diverges from related clades of the Bacillus cereus group in 16S-23S ribosomal DNA intergenic transcribed spacers containing tRNA genes. Appl Environ Microbiol 69:33–40. Navas M, Pincus DH, Wilkey K, Sercia L, Lasalvia M, Wilson D, Procop GW, Richter SS. 2014. Identification of aerobic Gram-positive bacilli by use of Vitek MS. J Clin Microbiol 52:1274–1277. Bohme K, Fernandez-No IC, Pazos M, Gallardo JM, Barros-Velazquez J, Canas B, Calo-Mata P. 2013. Identification and classification of seafood-borne pathogenic and spoilage bacteria: 16S rRNA sequencing versus MALDI-TOF MS fingerprinting. Electrophoresis 34:877– 887. Fernandez-No IC, Bohme K, Diaz-Bao M, Cepeda A, Barros-Velazquez J, Calo-Mata P. 2013. Characterisation and profiling of Bacillus subtilis, Bacillus cereus and Bacillus licheniformis by MALDI-TOF mass fingerprinting. Food Microbiol 33:235–242. Goodacre R, Shann B, Gilbert RJ, Timmins EM, McGovern AC, Alsberg BK, Kell DB, Logan NA. 2000.

136.

137.

138.

139.

140.

141.

142.

143.

144.

145.

146.

147.

148.

Detection of the dipicolinic acid biomarker in Bacillus spores using Curie-point pyrolysis mass spectrometry and Fourier transform infrared spectroscopy. Anal Chem 72:119–127. Le Fleche P, Hauck Y, Onteniente L, Prieur A, Denoeud F, Ramisse V, Sylvestre P, Benson G, Ramisse F, Vergnaud G. 2001. A tandem repeats database for bacterial genomes: application to the genotyping of Yersinia pestis and Bacillus anthracis. BMC Microbiol 1:2. Lista F, Faggioni G, Valjevac S, Ciammaruconi A, Vaissaire J, le Doujet C, Gorge O, De Santis R, Carattoli A, Ciervo A, Fasanella A, Orsini F, D’Amelio R, Pourcel C, Cassone A, Vergnaud G. 2006. Genotyping of Bacillus anthracis strains based on automated capillary 25loci multiple locus variable-number tandem repeats analysis. BMC Microbiol 6:33. Van Ert MN, Hofstadler SA, Jiang Y, Busch JD, Wagner DM, Drader JJ, Ecker DJ, Hannis JC, Huynh LY, Schupp JM, Simonson TS, Keim P. 2004. Mass spectrometry provides accurate characterization of two genetic marker types in Bacillus anthracis. Biotechniques 37:642– 644, 646, 648. Keim P, Price LB, Klevytska AM, Smith KL, Schupp JM, Okinaka R, Jackson PJ, Hugh-Jones ME. 2000. Multiple-locus variable-number tandem repeat analysis reveals genetic relationships within Bacillus anthracis. J Bacteriol 182:2928–2936. Priest FG, Barker M, Baillie LW, Holmes EC, Maiden MC. 2004. Population structure and evolution of the Bacillus cereus group. J Bacteriol 186:7959–7970. Tourasse NJ, Helgason E, Okstad OA, Hegna IK, Kolsto AB. 2006. The Bacillus cereus group: novel aspects of population structure and genome dynamics. J Appl Microbiol 101:579–593. Tourasse NJ, Okstad OA, Kolsto AB. 2010. HyperCAT: an extension of the SuperCAT database for global multischeme and multi-datatype phylogenetic analysis of the Bacillus cereus group population. Database (Oxford) 2010:baq017. Guinebretière MH, Auger S, Galleron N, Contzen M, De Sarrau B, De Buyser ML, Lamberet G, Fagerlund A, Granum PE, Lereclus D, De Vos P, Nguyen-The C, Sorokin A. 2010. Bacillus cytotoxicus sp. is a novel thermotolerant species of the Bacillus cereus group occasionally associated with food poisoning. Int J Syst Evol Microbiol 63:31–40. Apetroaie C, Andersson MA, Sproer C, Tsitko I, Shaheen R, Jaaskelainen EL, Wijnands LM, Heikkila R, Salkinoja-Salonen MS. 2005. Cereulide-producing strains of Bacillus cereus show diversity. Arch Microbiol 184:141–151. Carlin F, Fricker M, Pielaat A, Heisterkamp S, Shaheen R, Salonen MS, Svensson B, Nguyen-the C, EhlingSchulz M. 2006. Emetic toxin-producing strains of Bacillus cereus show distinct characteristics within the Bacillus cereus group. Int J Food Microbiol 109:132–138. Vassileva M, Torii K, Oshimoto M, Okamoto A, Agata N, Yamada K, Hasegawa T, Ohta M. 2007. A new phylogenetic cluster of cereulide-producing Bacillus cereus strains. J Clin Microbiol 45:1274–1277. Price EP, Seymour ML, Sarovich DS, Latham J, Wolken SR, Mason J, Vincent G, Drees KP, Beckstrom-Sternberg SM, Phillippy AM, Koren S, Okinaka RT, Chung WK, Schupp JM, Wagner DM, Vipond R, Foster JT, Bergman NH, Burans J, Pearson T, Brooks T, Keim P. 2012. Molecular epidemiologic investigation of an anthrax outbreak among heroin users, Europe. Emerg Infect Dis 18:1307–1313. Zwick ME, Joseph SJ, Didelot X, Chen PE, BishopLilly KA, Stewart AC, Willner K, Nolan N, Lentz S,

26. Bacillus and Other Aerobic Endospore-Forming Bacteria n

149.

150.

151.

152.

153.

154.

155. 156.

157.

158.

159.

Thomason MK, Sozhamannan S, Mateczun AJ, Du L, Read TD. 2012. Genomic characterization of the Bacillus cereus sensu lato species: backdrop to the evolution of Bacillus anthracis. Genome Res 22:1512–1524. Kuroda M, Serizawa M, Okutani A, Sekizuka T, Banno S, Inoue S. 2010. Genome-wide single nucleotide polymorphism typing method for identification of Bacillus anthracis species and strains among B. cereus group species. J Clin Microbiol 48:2821–2829. Pilo P, Frey J. 2011. Bacillus anthracis: molecular taxonomy, population genetics, phylogeny and patho-evolution. Infect Genet Evol 11:1218–1224. Van Ert MN, Easterday WR, Huynh LY, Okinaka RT, Hugh-Jones ME, Ravel J, Zanecki SR, Pearson T, Simonson TS, U’Ren JM, Kachur SM, Leadem-Dougherty RR, Rhoton SD, Zinser G, Farlow J, Coker PR, Smith KL, Wang B, Kenefic LJ, Fraser-Liggett CM, Wagner DM, Keim P. 2007. Global genetic population structure of Bacillus anthracis. PLoS One 2:e461. Borriss R, Chen XH, Rueckert C, Blom J, Becker A, Baumgarth B, Fan B, Pukall R, Schumann P, Sproer C, Junge H, Vater J, Puhler A, Klenk HP. 2011. Relationship of Bacillus amyloliquefaciens clades associated with strains DSM 7T and FZB42T: a proposal for Bacillus amyloliquefaciens subsp. amyloliquefaciens subsp. nov. and Bacillus amyloliquefaciens subsp. plantarum subsp. nov. based on complete genome sequence comparisons. Int J Syst Evol Microbiol 61:1786–1801. Quinn CP, Semenova VA, Elie CM, Romero-Steiner S, Greene C, Li H, Stamey K, Steward-Clark E, Schmidt DS, Mothershed E, Pruckler J, Schwartz S, Benson RF, Helsel LO, Holder PF, Johnson SE, Kellum M, Messmer T, Thacker WL, Besser L, Plikaytis BD, Taylor TH Jr., Freeman AE, Wallace KJ, Dull P, Sejvar J, Bruce E, Moreno R, Schuchat A, Lingappa JR, Martin SK, Walls J, Bronsdon M, Carlone GM, Bajani-Ari M, Ashford DA, Stephens DS, Perkins BA. 2002. Specific, sensitive, and quantitative enzyme-linked immunosorbent assay for human immunoglobulin G antibodies to anthrax toxin protective antigen. Emerg Infect Dis 8:1103–1110. Shlyakhov E, Shoenfeld Y, Gilburd B, Rubinstein E. 2004. Evaluation of Bacillus anthracis extractable antigen for testing anthrax immunity. Clin Microbiol Infect 10:421–424. Shlyakhov EN, Rubinstein E. 1994. Human live anthrax vaccine in the former USSR. Vaccine 12:727–730. Clinical and Laboratory Standards Institute. 2010. Methods for Antimicrobial Dilution and Disk Susceptibility Testing of Infrequently Isolated or Fastidious Bacteria; Approved Guideline, 2nd ed. CLSI document M45-A2. Clinical and Laboratory Standards Institute, Wayne, PA. Coker PR, Smith KL, Hugh-Jones ME. 2002. Antimicrobial susceptibilities of diverse Bacillus anthracis isolates. Antimicrob Agents Chemother 46:3843–3845. Mohammed MJ, Marston CK, Popovic T, Weyant RS, Tenover FC. 2002. Antimicrobial susceptibility testing of Bacillus anthracis: comparison of results obtained by using the National Committee for Clinical Laboratory Standards broth microdilution reference and Etest agar gradient diffusion methods. J Clin Microbiol 40:1902– 1907. Turnbull PC, Sirianni NM, LeBron CI, Samaan MN, Sutton FN, Reyes AE, Peruski LF Jr. 2004. MICs of selected antibiotics for Bacillus anthracis, Bacillus cereus, Bacillus thuringiensis, and Bacillus mycoides from a range

160.

161.

162.

163.

164.

165.

166.

167.

168.

169.

170.

171.

172.

173.

461

of clinical and environmental sources as determined by the Etest. J Clin Microbiol 42:3626–3634. Luna VA, King DS, Gulledge J, Cannons AC, Amuso PT, Cattani J. 2007. Susceptibility of Bacillus anthracis, Bacillus cereus, Bacillus mycoides, Bacillus pseudomycoides and Bacillus thuringiensis to 24 antimicrobials using Sensititre automated microbroth dilution and Etest agar gradient diffusion methods. J Antimicrob Chemother 60:555– 567. Centers for Disease Control and Prevention. 2001. Update: investigation of bioterrorism-related anthrax and interim guidelines for exposure management and antimicrobial therapy, October 2001. MMWR Morb Mortal Wkly Rep 50:909–919. Drago L. 2002. Bactericidal activity of levofloxacin, gatifloxacin, penicillin, meropenem and rokitamycin against Bacillus anthracis clinical isolates. J Antimicrob Chemother 50:1059–1063. Esel D, Doganay M, Sumerkan B. 2003. Antimicrobial susceptibilities of 40 isolates of Bacillus anthracis isolated in Turkey. Int J Antimicrob Agents 22:70–72. Weissfeld A, Snyder JW. 2013. Sentinel Level Clinical Laboratory Guidelines for Suspected Agents of Bioterrorism and Emerging Infectious Diseases: Bacillus anthracis. American Society for Microbiology, Washington, DC. http:// www.asm.org/index.php/guidelines/sentinel-guidelines. Stern EJ, Uhde KB, Shadomy SV, Messonnier NE. 2008. Conference report on public health and clinical guidelines for anthrax. Emerg Infect Dis 14:e1. Hendricks KA, Wright ME, Shadomy SV, Bradley JS, Morrow MG, Pavia AT, Rubinstein E, Holty JE, Messonnier NE, Smith TL, Pesik N, Treadwell TA, Bower WA, Workgroup on Anthrax Clinical Guidelines. 2014. Centers for Disease Control and Prevention expert panel meetings on prevention and treatment of anthrax in adults. Emerg Infect Dis 20:e130687. Meaney-Delman D, Zotti ME, Creanga AA, Misegades LK, Wako E, Treadwell TA, Messonnier NE, Jamieson DJ, Workgroup on Anthrax in Pregnant and Postpartum Women. 2014. Special considerations for prophylaxis for and treatment of anthrax in pregnant and postpartum women. Emerg Infect Dis 20:e130611. Weber DJ, Saviteer SM, Rutala WA, Thomann CA. 1988. In vitro susceptibility of Bacillus spp. to selected antimicrobial agents. Antimicrob Agents Chemother 32:642–645. Citron DM, Appleman MD. 2006. In vitro activities of daptomycin, ciprofloxacin, and other antimicrobial agents against the cells and spores of clinical isolates of Bacillus species. J Clin Microbiol 44:3814–3818. Logan NA, Hoffmaster AR, Shadomy SV, Stauffer KE. 2011. Bacillus and other aerobic endospore-forming bacteria, p 381–402. In Versalovic J, Carrol K, Funke G, Jorgensen JH, Landry ML, Warnock DW (ed), Manual of Clinical Microbiology, 10th ed. ASM Press, Washington, DC. O’Day DM, Smith RS, Gregg CR, Turnbull PC, Head WS, Ives JA, Ho PC. 1981. The problem of Bacillus species infection with special emphasis on the virulence of Bacillus cereus. Ophthalmology 88:833–838. Ligozzi M, Lo Cascio G, Fontana R. 1998. vanA gene cluster in a vancomycin-resistant clinical isolate of Bacillus circulans. Antimicrob Agents Chemother 42:2055–2059. Patel R, Piper K, Cockerill FR III, Steckelberg JM, Yousten AA. 2000. The biopesticide Paenibacillus popilliae has a vancomycin resistance gene cluster homologous to the enterococcal VanA vancomycin resistance gene cluster. Antimicrob Agents Chemother 44:705–709.

Listeria and Erysipelothrix NELE WELLINGHAUSEN

27 exceptionally large growth temperature range from 0 to 50°C. The optimum growth temperature is between 30 and 37°C, but at 4°C, growth is also observed within a few days. Catalase is typically produced, but catalase-negative strains causing disease in humans have been described (8, 9). The oxidase test is negative. Acid is produced from D-glucose and other sugars. The Voges-Proskauer and methyl red tests are positive. Esculin is hydrolyzed in a few hours. Urea and gelatin are not hydrolyzed. Neither indole nor H2S is produced. The cell wall contains a directly cross-linked peptidoglycan based on meso-diaminopimelic acid, as well as lipoteichoic acid, but no mycolic acids. The two predominant cellular fatty acids are Cai15:0 and Cai17:0 (branchedchain type) (10).

LISTERIA Taxonomy The genus Listeria consists of Gram-positive, non-sporeforming, facultative anaerobic, regular rod-shaped bacteria with a low G+C content of 36 to 42 mol%. While early phylogenetic studies suggested a close relation between Listeria and the Lactobacillaceae, comparisons of 16S rRNA gene sequences have shown that Listeria is most closely related to Staphylococcus and Bacillus. Listeria belongs to the newly assigned family Listeriaceae within the order Bacillales (1). Synthesis of menaquinones and major amounts of branched-chain fatty acids confirms the taxonomic separation of Listeria from the Lactobacillaceae (2). Until recently, the genus Listeria comprised six validated species including Listeria monocytogenes as the type species in the genus, L. grayi, L. innocua, L. ivanovii, L. seeligeri, and L. welshimeri. Within L. ivanovii, two subspecies, L. ivanovii subsp. ivanovii and L. ivanovii subsp. londoniensis, are differentiated. In recent years, further Listeria species, i.e., L. marthii, L. rocourtiae, L. fleischmannii, including a recent subspecies L. fleischmannii subsp. coloradonensis (3), and L. weihenstephanensis, have been described from the natural environment and from food items (4–7). Based on the results of multilocus enzyme electrophoresis, DNA-DNA hybridization, and 16S rRNA gene sequencing, a group of species closely related to L. monocytogenes, including L. innocua, L. ivanovii, L. marthii, L. monocytogenes, L. seeligeri, and L. welshimeri, can be differentiated from L. grayi and the remaining recently described species (4–7). Only L. monocytogenes and L. ivanovii are pathogenic for humans and animals.

Epidemiology and Transmission The primary habitat of Listeria species is the environment, where they exhibit a saprophytic lifestyle. L. monocytogenes has been isolated from various animals, like mammals, birds, fish, and crustaceans. Infected animals can asymptomatically pass the organism or develop clinical disease. Due to its widespread distribution, L. monocytogenes has many opportunities to enter human food production, resulting in contamination of fresh and processed poultry, meat, and vegetables; raw milk; cheese; smoked salmon; etc. Numbers of organisms exceeding 103 CFU/g were detected in food products (11). Infection of humans ingesting colonized food is potentiated by the ability of the organism to multiply at 4°C. The intestinal tract of adults is consistently colonized with nonpathogenic Listeria species and, to a lesser extent (1 to 5%), with pathogenic L. monocytogenes. Cervicovaginal carriage in women has not been reported. Apart from foodrelated infections, nosocomial outbreaks, mainly in neonatal wards, have been described (12, 13). The number of sporadic cases of listeriosis in countries that report the illness is typically in the range of 0.1 to 0.9 cases per 100,000 persons. While the number of cases and the mortality in the United States have decreased until the end of the last millennium, the incidence remained quite stable in the last decade around 0.27 cases per 100,000 persons (14–17). In contrast, in several European countries, the incidence of sporadic cases of listeriosis has increased, reaching numbers from 0.4 up to 1.0 per 100,000 per year (18–20). People with an underlying condition, like chronic lymphocytic leukemia, myeloproliferative disorder, cancer, organ transplantation, alcoholism, and hepatic disorders;

Description of the Agent Members of the genus Listeria are Gram-positive, facultative anaerobic, non-spore-forming, nonbranching, regular, short (0.5 to 2 μm by 0.4 to 0.5 μm) rods that occur singly or in short chains. Filaments of 6 to 20 μm in length may occur in older or rough cultures. Temperature-regulated expression of flagellin results in a characteristic tumbling motility at 20 to 28°C by means of one to six peritrichous flagella. At 37°C, the organisms are much less motile. Colonies are small (1 to 2 mm in diameter after 1 or 2 days of incubation at 37°C), smooth, and blue-gray on nutrient agar when examined with obliquely transmitted light and typically show a narrow zone of beta-hemolysis. Listeria spp. show an

doi:10.1128/9781555817381.ch27

462

27. Listeria and Erysipelothrix

pregnant women; and individuals ≥60 years have an up to 100 to >1,000-fold increased risk of acquiring listeriosis (15, 17, 18, 20).

Clinical Significance In adults, L. monocytogenes causes primarily septicemia, meningitis, and encephalitis with a mortality reaching up to 50%. According to a recent study, the rate of unfavorable outcome of Listeria meningitis has increased significantly during the last 15 years and could be linked to infection with L. monocytogenes serotype 6 (21). Focal infections with Listeria spp. have been infrequently described and include endocarditis, pericarditis, arthritis, osteomyelitis, intra-abdominal and brain abscesses, endophthalmitis, (sclero-)keratitis, peritonitis, cholecystitis, and intravenous catheter and pleuropulmonary infections (22–24). Among veterinarians and abattoir workers, but recently also in a gardener, primary cutaneous listeriosis with or without bacteremia has been reported (25, 26). In pregnant women, L. monocytogenes often causes a mild, self-limited influenza-like illness. Transient bacteremia can result in placentitis and/or amnionitis, and since Listeria is able to cross the placenta, it can infect the fetus, causing abortion, stillbirth, or most commonly, preterm labor (27, 28). In neonates, an early-onset form and a lateonset form of listeriosis occur. The early form is presumably caused by intrauterine infection and manifests as granulomatosis infantisepticum. The organism is widely disseminated in the body, including the central nervous system. The source of the organism in the late-onset cases, which manifest at a mean of 14 days after birth, is unclear and may comprise the mother’s genital tract or environmental sources. The incubation period for human listeriosis varies between 1 day and 2 to 3 months and is significantly longer for pregnancy-associated cases (median, 28 days) than for central nervous system and bacteremia cases (median of 9 and 2 days, respectively) (29). The infectious dose has not been firmly established, but 105 CFU or greater have been reported to cause gastroenteritis in outbreak situations (30). A dose-response model using rhesus monkeys as a surrogate for pregnant women recently indicated that oral exposure to 107 CFU of L. monocytogenes results in about 50% stillbirths (31). In a pregnant guinea pig model, doses of 104 to 108 CFU have been shown to invade fetal liver and brain tissue in up to 75%, and fetal infection occurred as early as day 2 after maternal infection (32). Thus, the infectious dose may be much less than the extrapolated estimate of 1013 CFU from the FDA-U.S. Department of Agriculture-CDC risk assessment based on mouse data (33). Most cases of Listeria gastroenteritis are linked to foodborne outbreaks (34). Typically, patients with Listeria gastroenteritis have no known predisposing risk factors for listeriosis, illness occurs about 24 h after ingestion of a food item that is contaminated with a large number of bacteria (105 to 109 CFU/g or ml), and illness lasts about 2 days. Apart from gastroenteritis, fever, headache, and pain in joints and muscles are frequently seen. After ingestion of L. monocytogenes, pathogen and host factors as well as the number of pathogens ingested determine whether invasive infection develops. Immunity to listeriosis is effected primarily via the cell-mediated immune system. Penetration of the epithelial barrier in the gut by L. monocytogenes is facilitated by its ability to escape from the host cell vacuole, intracytoplasmic multiplication, movement via bacterially induced polymerization of host cell actin, and spread to neighboring cells through pseudo-

n

463

pod-like extensions of the host cell membrane. By N-deacetylation of peptidoglycan of the cell wall, Listeria evades innate immune defenses (35). Virulence genes are clustered on an 8.2-kb pathogenicity island and include genes coding for internalin A and B and listeriolysin, a hemolysin (36). Interaction between internalin and E-cadherin, a receptor of the trophoblast, facilitates the spread of the organism to the fetus. L. ivanovii is primarily a pathogen of ruminants, but systemic infections in humans with underlying conditions and a case of stillbirth in a pregnant woman have been described (37).

Collection, Transport, and Storage of Specimens Suitable specimens for detection of listeriosis include blood and cerebrospinal fluid (CSF). In neonates with suspicion of listeriosis, investigation of blood, CSF, amniotic fluid, respiratory secretions, placental or cutaneous swabs, gastric aspirates, or meconium can facilitate detection of the organism. For epidemiologic purposes or rare causes of gastroenteritis, stool specimens are preferred to rectal swabs. In general, specimens for detection of Listeria do not need special handling during collection. Clinical specimens for culture of L. monocytogenes should be processed as soon as possible or stored and transported at room temperature or 4°C for up to 48 h. At 4°C, even longer storage times may be tolerated due to the specific cold resistance of the organism, but multiplication of Listeria has to be regarded. Stool samples (1 g each) can be inoculated into 100 ml of a selective broth, e.g., University of Vermont, polymyxin-acriflavine-lithium chloride-ceftazidime esculin-mannitol (PALCAM), or Listeria enrichment broth (38–41), analogous to the preparation of food samples (see below), and then shipped overnight at room temperature. Nevertheless, the value of enrichment of clinical samples has not been investigated yet. To avoid overgrowth of L. monocytogenes by contaminating microbiota during longer periods of storage, nonsterile-site specimens should be stored at 4°C for 24 to 48 h or frozen at −20°C. Food samples should include a minimum of 100 g of a sample and should be collected aseptically in sterile containers. Food packaged in original containers should always be preferred. Samples should be shipped overnight frozen. Although L. monocytogenes is relatively resistant to freezing, repeated freezing and thawing should be avoided. Cultures of Listeria spp. should be frozen at −20 to −70°C for long-term storage. They can be shipped on a non-glucose-containing agar slant and packaged and declared according to the respective national and international requirements. Because L. monocytogenes can infect the fetus, pregnant women should be particularly careful when working in a laboratory where L. monocytogenes is propagated or handled.

Direct Examination Direct microscopy should be performed on CSF, positive blood cultures, and if available, tissue samples. Detection of Gram-positive, regular short rods in CSF or blood cultures should lead to the suspicion of listeriosis. Nevertheless, L. monocytogenes may be confused with members of the coryneform rods (especially in direct slides from positive blood cultures), since the cells may be arranged in V forms or palisades. Commercial tests licensed for antigen detection in clinical specimens are not available.

464 n BACTERIOLOGY

Sensitive and specific in-house PCR assays have been described for detection of L. monocytogenes in CSF, stool, or lung tissue (39, 42–46) and may be particularly useful for specimens from patients with prior antimicrobial therapy. Regarding commercial assays, the Probelia Listeria monocytogenes assay (Bio-Rad, Hercules, CA) has been evaluated in clinical stool specimens, while other commercial assays (e.g., LightCycler PCR, Roche Diagnostics, Indianapolis, IN; TaqMan Listeria monocytogenes detection kit, Applied Biosystems, Carlsbad, CA; BAX System PCR, DuPont, Wilmington, DE) have been validated only for food specimens. Recently, a fluorescent in situ hybridization assay has been described that allows detection of L. monocytogenes in wastewater samples with a sensitivity of 104 cells/ml (47).

Isolation Procedures Clinical specimens from normally sterile sites should be plated onto tryptic soy agar containing 5% sheep, horse, or rabbit blood. Plates should be incubated at 35 to 37°C in an atmosphere enriched with 5% CO2 for a minimum of 48 h. Listeria colonies typically show a narrow zone of betahemolysis on blood agar. Blood samples should be inoculated into conventional blood culture media. Clinical specimens obtained from nonsterile sites, like stool samples, as well as food and environmental specimens should be plated on Listeria spp. selective agars. In addition, enrichment by inoculation into selective broth for Listeria spp. should be done before plating. Selective agars for culture of Listeria spp. include lithium chloride-phenylethanol-moxalactam (LPM) (48), Oxford, modified Oxford, and PALCAM agars (39). On LPM agar, colonies have to be examined under a stereomicroscope with Henry illumination (magnification, ×15 to ×25, with oblique lighting directed to the microscope stage by a concave mirror positioned at a 45° angle to the incident light). Listeria colonies appear blue, and colonies of other bacteria appear yellowish or orange. Oxford and PALCAM agars contain selective substances that eliminate the need for examination under oblique lighting. On Oxford and modified Oxford agars, Listeria colonies appear black due to esculin hydrolysis, are 1 to 3 mm in diameter, and are surrounded by a black halo after 24 to 48 h of incubation at 35 to 37°C. On PALCAM agar, Listeria colonies appear graygreen, are approximately 2 mm in diameter, and have black sunken centers. For the detection of Listeria spp. in food samples, enrichment methods have to be used. The most widely used reference methods for food and environmental samples are the Food and Drug Administration (FDA) Bacteriological and Analytical Manual (41) and the U.S. Department of Agriculture (USDA) method (40) in the United States and the International Organization for Standards (ISO) 11290 method in Europe (49). All methods require enrichment of the samples in a selective broth (buffered Listeria enrichment broth in the FDA Bacteriological and Analytical Manual method, University of Vermont broth in the USDA method, and Fraser broth in the ISO method) prior to plating onto selective agar (see above; also agar Listeria according to Ottaviani and Agosti [ALOA]) and biochemical identification of typical colonies (40, 41). Confirmation of species identification is necessary, since growth of Bacillus circulans strains on ALOA has been described (50). A detailed comparison of methods is given by Dever et al. (51). Chromogenic media allow selective isolation of Listeria species (39, 52, 53). Chromogenic media are mainly based on ALOA and include ALOAgar (Biolife, Milan, Italy), BCM L. monocytogenes (Biosynth, Staad, Switzerland), Li-

mono-Ident-Agar (Heipha, Eppelheim, Germany), Chromoplate Listeria (Merck KGaA, Darmstadt, Germany), Oxoid Chromogenic Listeria agar (Oxoid, Basingstoke, United Kingdom), and BBL Chromagar (BD, Sparks, MD) (53–55). However, none of these agars differentiate between L. monocytogenes and L. ivanovii. Specific detection of L. monocytogenes is facilitated on Rapid L.mono agar (Bio-Rad, Hercules, CA) (56). Chromogenic media showed sensitivities comparable to those of Oxford and PALCAM agar, but they appeared to be less specific (52, 53, 55, 56).

Identification A simplified identification is based on the following tests: Gram staining, observation of tumbling motility in a wet mount, and tests for a positive catalase reaction and esculin hydrolysis. Acid production from D-glucose and a positive Voges-Proskauer test are confirmatory results. Listeria spp. may be confused with other Gram-positive bacteria due to similar morphologic or biochemical characteristics. Streptococcus and Enterococcus spp. can be differentiated from Listeria spp. on the basis of Gram stain morphology, motility, and catalase reaction. Erysipelothrix spp. differ from Listeria spp. in motility, catalase reaction, and ability to grow at 4°C (Erysipelothrix spp. do not grow at that temperature). Lactobacillus spp. are usually nonmotile and catalase negative. Identification of Listeria isolates to the species level is crucial, because all species can contaminate foods but only L. monocytogenes is of public health concern. A scheme for identification of medically relevant Listeria species based on morphological and biochemical characteristics is shown in Table 1. Among these markers, hemolysis is essential for differentiating between L. monocytogenes and the most frequently isolated nonpathogenic species, L. innocua. Production of hemolysin is regarded as a key virulence factor of L. monocytogenes and is visible on sheep blood agar plates as a narrow zone of beta-hemolysis that frequently does not extend much beyond the edges of the colonies (Fig. 1). Like L. monocytogenes, L. ivanovii is also hemolytic, but hemolysis alone cannot be used to discriminate pathogenic and nonpathogenic species, since L. seeligeri is, besides rare exceptions (57), hemolytic and since hemolytic strains of L. innocua have been described as well (58). The CAMP (Christie, Atkins, Munch-Petersen) test can be used to differentiate among hemolytic Listeria species. The test is carried out by streaking a beta-hemolysisproducing Staphylococcus aureus strain and Rhodococcus equi parallel to each other on a blood agar plate. Suspect cultures are streaked at right angles in between (but not touching) the two streaks. Hemolysis by L. monocytogenes and, to a lesser degree, L. seeligeri is enhanced in the vicinity of S. aureus, and hemolysis by L. ivanovii is enhanced in the vicinity of the R. equi streak (Fig. 2). However, the reliability of the CAMP test is limited. As recommended in the USDA method (40), commercially available β-lysis disks (Remel, Lenexa, KS) may be used instead. L. monocytogenes is, apart from rare atypical strains, Lrhamnose and α-methyl-D-mannoside positive and D-xylose negative. Incubation of test tubes for up to 7 days (37°C, aerobically) may be necessary. Commercially available miniaturized tests considerably speed up biochemical identification of Listeria spp. The API Coryne and Listeria (bioMérieux, Durham, NC), Micro-ID Listeria (Remel), BBL Crystal Gram-Pos ID (BD, Franklin Lakes, NJ), and Microbact Listeria 12L (Oxoid) reliably identify Listeria isolates to the genus and species level (59).

n

27. Listeria and Erysipelothrix TABLE 1

465

Biochemical differentiation of medically relevant and selected environmental species in the genus Listeriaa

Characteristic

L. grayi

L. innocua

L. ivanovii subsp. ivanovii

L. ivanovii subsp. londoniensis

L. monocytogenes

L. seeligeri

L. welshimeri

Beta-hemolysis CAMPc test reaction S. aureus R. equi Acid production from: Mannitol α-Methyl-Dmannoside L-Rhamnose Soluble starch D-Xylose Ribose N-Acetyl-βD-mannosamine Hippurate hydrolysis Reduction of nitrate Associated serovar(s)

−

−

++b

++

+

+

−

− −

− −

− +

− +

+ V

+ −

− −

+ +

− −

− −

− −

− +

− −

− +

V + − V ND

V − − − ND

− − + + V

− − + − +

+ − − − ND

− ND + − ND

V ND + − ND

− V S

+ − 4ab, 6a, 6b

+ − 5

+ − 5

+ − 1/2a, 1/2b, 1/2c, 3a, 3b, 3c, 4a, 4ab, 4b, 4c, 4d, 4e, 7

ND ND 1/2a, 1/2b, 1/2c, 4a, 4b, 4d, 6b

ND ND 1/2b, 4c, 6a, 6b

a Symbols and abbreviations: +, ≥90% of strains are positive; −, ≥90% of strains are negative; ND, not determined; V, variable; US, undesignated serotype; S, specific. b ++, usually a wide zone or multiple zones. c See text and Fig. 2.

FIGURE 1 Macroscopic view of colonies on 5% human blood agar plates after 24 h of incubation. (A) L. monocytogenes: discrete zone of beta-hemolysis under the removed colonies. (B) L. innocua: no hemolysis. (C) L. ivanovii: wide zone of beta-hemolysis around the colonies. doi:10.1128/9781555817381.ch27.f1

466 n

BACTERIOLOGY

Billerica, MA; Vitek MS, bioMérieux; in-house databases) and an optional pretreatment protocol for colonies (64– 66). The use of MALDI-TOF (MS) on the Andromas system directly from colony material facilitated species identification of L. grayi, while L. monocytogenes clustered with all other Listeria species and was, thus, identifiable to the genus level only. By use of the Vitek MS v2.0 system, 75% of the tested L. monocytogenes isolates could be identified to the species level and 9% to the genus level, while 15% failed to be identified (66). Differentiation of L. grayi from a cluster including L. monocytogenes and all other Listeria species by MALDI-TOF (MS) corroborates molecular and phylogenetic studies of the genus Listeria (67).

Typing Systems

FIGURE 2 CAMP test done with S. aureus CIP 5710 (top plate) and R. equi CIP 5869 (bottom plate) after 24 h of incubation. Upper right, L. monocytogenes; lower right, L. innocua; middle left, L. ivanovii. doi:10.1128/9781555817381.ch27.f2

The RapID CB Plus system (Oxoid) allows reliable identification of Listeria species to the genus level only (60, 61). Identification of L. monocytogenes is possible with the Vitek2 (bioMérieux), MicroScan WalkAway (Siemens Healthcare, Malvern, PA), Micronaut (Merlin, BornheimHesel, Germany), Phoenix (BD), and Biolog (Biolog, Inc., Hayward, CA) systems. All systems allow reliable species identification, while Vitek1 (bioMérieux) facilitates identification of Listeria isolates to the genus level only. A chemiluminescence DNA probe assay (AccuProbe, Hologic Gen-Probe, Bedford, MA) (62) is available for rapid identification of L. monocytogenes from primary isolation plates. However, false-positive results were observed with L. marthii (5). Recently described peptide nucleic acid fluorescent in situ hybridization probes also allow rapid and specific identification of L. monocytogenes and L. ivanovii (63). The matrix-assisted laser desorption ionization–time of flight (mass spectrometry) (MALDI-TOF [MS]) technique is increasingly used, especially in European laboratories, and allows discrimination of the Listeria species by use of the respective software (MALDI Biotyper, Bruker Daltonics,

Subtyping of L. monocytogenes is crucial for the workup of disease acquired from food-borne agents. Based on somatic “O” and flagellar “H” antigens, 13 serovars of L. monocytogenes are known (1/2a, 1/2b, 1/2c, 3a, 3b, 3c, 4a, 4ab, 4b/4bX, 4c, 4d, 4e, and 7). Since the vast majority of L. monocytogenes strains that cause sporadic infections or outbreaks belong to the same serotypes, i.e., 1/2a, 1/2b, and 4b, and since serotyping antigens are shared among L. monocytogenes, L. innocua, L. seeligeri, and L. welshimeri, reliable discrimination below the level of serotype is necessary. Thus, serotyping is only useful as a first-level discriminator or for the selection of further typing methods in suspected outbreaks. Antisera are commercially available from Difco (Difco Laboratories/BD, Sparks, MD) and Denka Seiken (Tokyo, Japan). Multiplex PCR assays, also validated by interlaboratory comparison, have been described for identification of the four major serovars of L. monocytogenes (1/2a, 1/2b, 1/2c, and 4b) (68–72). Pulsed-field gel electrophoresis (PFGE) is considered the standard typing method for L. monocytogenes and is particularly useful for subtyping of serovar 4b isolates. Its discriminatory power and reproducibility of results have been confirmed in a World Health Organization multicenter international typing study (73) as well as in a large number of other studies. To optimize quality and interlaboratory comparability of PFGE, standardized laboratory protocols should be used. PulseNet USA has published a 1-day protocol (74) (available at http://www.cdc.gov/ pulsenet/PDF/listeria-pfge-protocol-508c.pdf) that has recently been optimized and re-evaluated by a multilaboratory study (75). According to the protocol, restriction endonucleases ApaI and AscI are used. A proficiency testing trial of the PulseNet Europe involving 29 national reference laboratories showed a high level of agreement between typing results but also stressed the necessity of adherence to the standardized protocol and revealed failures in DNA extraction (72). Faster and simpler molecular subtyping methods, like multilocus variable-number tandem-repeat analysis and multilocus sequence typing, have evolved, and their application for subtyping of L. monocytogenes is supported by PulseNet. Both methods showed a discriminatory power comparable to that of PFGE (76–79). By multilocus sequence typing and multi-virulence locus sequence typing, the predominant PFGE clone of L. monocytogenes involved in recent listeriosis outbreaks in Canada could be further characterized and traced back to 1988 (80). Other molecular typing methods, like single-nucleotide polymorphism-based multilocus genotyping and a mixed-genome DNA microarray, have been developed and also showed results comparable to those of PFGE (81–84). High-throughput genome pyrosequencing allowed rapid molecular characterization of the outbreak strain of L. monocytogenes serotype 1/2a in a

27. Listeria and Erysipelothrix

large listeriosis outbreak in Canada in 2008, confirming differences in AscI PFGE patterns at the molecular level (85). Altogether, sequence-based molecular methods may further improve subtyping of L. monocytogenes. They may replace typing methods with high discriminatory power but lacking interlaboratory standardization, like random amplification of polymorphic DNA (RAPD), amplified fragment length polymorphism, multilocus single-strand conformation polymorphism, and ribotyping, and may also add relevant molecular information about outbreak strains to the knowledge obtained by PFGE. As a nonmolecular technique, MALDI-TOF (MS) has been used successfully for rapid typing of L. monocytogenes. It allowed clear discrimination of all lineages and serotypes of L. monocytogenes in one study so far (64). An overview of the methods used for typing of L. monocytogenes is given by Jadhav et al. (44).

Serologic Tests Antibodies directed against listeriolysin O have been detected in listeriosis patients by blotting techniques with sensitivities from 50 to 96%, but the sensitivity was markedly lower with complement fixation or O-agglutination tests (86, 87). A test based on the detection of antibodies against recombinant truncated forms of listeriolysin O may be more specific (88). Serologic tests cannot be recommended for the detection of past or acute listeriosis.

n

467

normally sterile sites. Species identification is necessary to differentiate L. monocytogenes from nonpathogenic Listeria species. Especially for patients with underlying immunosuppression and for individuals older than 60 years and younger than 1 month, direct microscopic detection of Gram-positive, regular, short rods in the above-mentioned specimens should raise suspicion of listeriosis and should promptly be communicated to the clinician to ensure eradication of L. monocytogenes by antimicrobial therapy. Detection of Listeria species in stool samples likely represents colonization. Routine screening of stool samples for Listeria remains unwarranted, although sporadic cases of L. monocytogenes gastroenteritis have been reported. Standard antimicrobial therapy of meningitis with cefotaxime or ceftriaxone is not active against L. monocytogenes. Antimicrobial susceptibility testing should be performed in cases of suspected treatment failures, severe disease, and patients with penicillin allergy. Cultures from blood and CSF that were obtained after the initiation of antimicrobial therapy may be negative. In these cases, detection of Listeria DNA may be useful. Commercial kits for PCR-based detection of L. monocytogenes in CSF specimens are not available, but in-house protocols and multiplex PCR formats facilitate rapid and specific detection of L. monocytogenes DNA.

ERYSIPELOTHRIX

Antimicrobial Susceptibilities

Taxonomy

Treatment with an aminopenicillin (ampicillin or amoxicillin) is still regarded as the most effective therapeutic regimen for listeriosis. In vitro resistance of L. monocytogenes isolates to ampicillin has been reported sporadically (89–91), but the methods used in these studies appear to be inadequate for susceptibility testing of Listeria. Therefore, these reports have to be interpreted with great caution. In other studies and reviews, in vitro resistance against ampicillin was not detected (92–94). Since aminoglycosides exhibit a synergistic effect on aminopenicillins, they are usually added for therapy of listeriosis. However, in a retrospective cohort study, addition of gentamicin did not decrease early or late mortality in listeriosis (95). Trimethoprimsulfamethoxazole is recommended for patients who are allergic to penicillin, and moxifloxacin may be a valuable alternative, since it shows bactericidal efficacy comparable to that of amoxicillin in vitro (96). L. monocytogenes is intrinsically resistant to cephalosporins, fosfomycin, and fusidic acid, and even when in vitro susceptibility may be determined, cephalosporins should not be used for therapy. Isolated resistance against tetracycline has been noted (97) as well as multidrug resistance to chloramphenicol, macrolides, and tetracyclines due to the presence of resistance plasmids. Newer substances against Gram-positive pathogens, like linezolid, tigecycline, and daptomycin, elicited high susceptibility in vitro, and linezolid also showed clinical efficacy (98– 102). In addition, L. monocytogenes is generally susceptible in vitro to erythromycin and vancomycin (97, 103). Antimicrobial susceptibility testing should be performed in cases of suspected treatment failures, severe disease, and patients with penicillin allergy. A CLSI guideline (M45-A2) for broth microdilution antimicrobial susceptibility testing of L. monocytogenes including interpretive breakpoints for penicillin, ampicillin, and trimethoprim-sulfamethoxazole is available (104).

Erysipelothrix is taxonomically classified within the class Erysipelotrichia (105) in the phylum Firmicutes. The genus Erysipelothrix has three validly published species, E. rhusiopathiae, E. tonsillarum, and E. inopinata. Based on peptidoglycan antigens of the cell wall, several serovars can be distinguished in E. rhusiopathiae (serovars 1a, 1b, 2a, 2b, 3, 4, 5, 6, 8, 9, 11, 12, 15, 16, 17, 19, 21) and E. tonsillarum (serovars 3, 7, 10, 14, 20, 22, and 23) (106). Only E. rhusiopathiae has been detected as a pathogen of humans. The vast majority of infections are caused by serovars 1 and 2.

Evaluation, Interpretation, and Reporting of Results The diagnosis of listeriosis can be made by isolation of L. monocytogenes from blood, CSF, or specimens from other

Description of the Agent Erysipelothrix organisms are facultatively anaerobic, nonspore-forming, non-acid-fast, Gram-positive bacteria that appear microscopically as short rods (0.2 to 0.5 μm by 0.8 to 2.5 μm) with rounded ends and occur singly, in short chains, or in long, nonbranching filaments (60 μm or more in length). Some cells stain unevenly. They are nonmotile and grow in complex media at a wide range of temperatures (5 to 42°C; optimum, 30 to 37°C) and at alkaline pH (pH 6.7 to 9.2; optimum, pH 7.2 to 7.6). Like Listeria organisms, they can grow in the presence of high concentrations of sodium chloride (up to 8.5%). Erysipelothrix organisms are catalase negative and oxidase negative, do not hydrolyze esculin, and weakly ferment glucose without the production of gas. They are methyl red and Voges-Proskauer negative and do not produce indole or hydrolyze urea but distinctively produce H2S in triple sugar iron agar. Key fatty acids are C16:0 and C18:cis9 (10).

Epidemiology and Transmission E. rhusiopathiae is distributed worldwide in nature and is remarkably stable under varying environmental conditions. The organism is carried by a variety of animals, like mammals, birds, and fish, in their digestive tracts or tonsils but is most frequently associated with pigs. Other domestic animals that are frequently infected include sheep, rabbits, cattle, and turkeys. Infected animals, both sick and asymp-

468 n

BACTERIOLOGY

tomatic, pass the organism by urine and feces, leading to contamination of water and soil. Infection in animals is most likely acquired by ingestion of contaminated matter. Human infection with E. rhusiopathiae is a zoonosis. Most cases are related to occupational exposure, occurring most frequently among fish handlers, veterinarians, and butchers. The disease is contracted through direct contact via skin abrasions, injuries, or animal bites (107). E. tonsillarum has been recovered from tonsils of healthy pigs and cattle, water, and seafood. E. inopinata has been isolated once from a vegetable-based peptone broth (108).

Clinical Significance E. rhusiopathiae has been recognized for more than 100 years as the agent of swine erysipelas, an acute or chronic disease. In humans, E. rhusiopathiae causes erysipeloid, a localized cellulitis developing within 2 to 7 days around the inoculation site. The infected area is swollen, and the mostly painful lesion consists of a well-defined, slightly elevated, violaceous zone which spreads peripherally as discoloration of the central area fades. Vesicles may be present, but suppuration does not occur. Regional lymphangitis is present in onethird of patients, and low-grade fever and arthralgias occur in about 10% of patients. Healing of erysipeloids usually takes 2 to 4 weeks and sometimes months, and relapses are frequently seen. Dissemination of the organism can occur and manifests in most of the cases as endocarditis with a poor prognosis (107). Uncommon manifestations of infection with E. rhusiopathiae include peritonitis, endophthalmitis, osteomyelitis, intracranial and spinal abscesses, prosthetic joint arthritis, pneumonia, and meningitis (109–111). Data about E. rhusiopathiae pathogenesis are still scarce. E. rhusiopathiae has a capsule consisting of polysaccharide antigen that confers increased resistance to phagocytosis. Neuraminidase plays a significant role in bacterial attachment and subsequent invasion into host cells. The 69-kDa surface antigens SpaA, SpaB, and SpaC appear to be the major protective antigens of E. rhusiopathiae, and recombinant SpaA and SpaC elicit a protective immune response in pigs and mice, making them potential vaccine candidates (112, 113). A multiplex real-time PCR for detection of Spa types has been described (114).

Collection, Transport, and Storage of Specimens Biopsy specimens from erysipeloid lesions are the best source of E. rhusiopathiae. Care should be taken to cleanse and disinfect the skin before sampling. The organisms typically are located deep in the subcutaneous layer of the leading edge of the lesion; hence, a biopsy of the entire thickness of the dermis at the periphery of the lesion should be taken for Gram staining and culture. Swabs from the surface of the skin are not useful. In disseminated disease, the organism can be cultured in standard blood cultures or from aspirates of the respective infected location. For transport and storage of specimens, standard procedures should be followed.

Direct Examination Direct microscopy should be performed with aspirates, biopsy specimens, and positive blood cultures. Gram stain morphology of E. rhusiopathiae includes short rods and very long filaments and, thus, is not distinctive. However, the presence of long, slender, Gram-positive rods in tissue from an individual with a known exposure is suggestive of erysipeloid. It has to be noted that the organism may appear Gram negative in stains from cultures (see below).

Conventional and real-time PCR assays for specific detection of E. rhusiopathiae in animal tissue as well as for discrimination of E. rhusiopathiae from E. tonsillarum have been described, but their application to human samples has not been evaluated yet (115–117).

Isolation Procedures Tissue or biopsy specimens should be processed as described in chapter 18 and plated onto blood agar or chocolate blood agar, placed in tryptic soy, Schaedler, or thioglycolate broth, and incubated at 35 to 37°C aerobically or in 5% CO2 for 7 days. Special pretreatment of samples is not necessary, but inoculation of an enrichment broth significantly increases the detection rate. Blood from patients with septicemia or endocarditis can be inoculated into commercial blood culture systems. E. rhusiopathiae colonies generally develop in 1 to 3 days, appearing as pinpoints (105/ml. The clinical significance of coryneform bacteria is strengthened by the following findings: (i) multiple specimens are positive for the same coryneform bacteria, (ii) coryneform bacteria are seen in the direct Gram stain and a strong leukocyte reaction is also observed, and (iii) other organisms recovered from the same material are of low pathogenicity. For a comprehensive summary of case reports on individual coryneform bacteria, the reader is referred to review articles (2, 47, 48). The most frequently reported coryneforms as well as their established disease associations are listed in Table 2. Historically, diphtheria caused by C. diphtheriae (or C. ulcerans) is the most prominent infectious disease for which coryneform bacteria are responsible. Therefore, special attention is given to that disease in this chapter; a comprehensive review on diphtheria has been recently published (49). Due to immunization programs, the disease has nearly disappeared in countries with high socioeconomic standards. However, the disease is still endemic in some subtropical and tropical countries as well as among individuals of certain

Most frequently reported disease associations of coryneform bacteria in humans

C. amycolatum

pseudodiphtheriticum pseudotuberculosis resistens riegelii striatum

C. tuberculostearicum C. ulcerans (toxigenic) C. urealyticum Arthrobacter spp. Brevibacterium spp. Dermabacter hominis Helcobacillus sp. Rothia spp. Cellulomonas spp. Cellulosimicrobium sp. Microbacterium spp. A. haemolyticum T. bernardiae T. pyogenes G. vaginalis

479

Disease or disease association Wound infections, foreign body infections, bacteremia, sepsis, urinary tract infections Genitourinary tract infections (mainly females) Urinary tract infections Throat diphtheria, cutaneous diphtheria Endocarditis, foreign body infections, pharyngitis Genitourinary tract infections (mainly males) Endocarditis, bacteremia, foreign body infections, wound infections Granulomatous lobular mastitis Eye infections Wound infections, urinary tract infections, respiratory tract infections Respiratory tract infections, endocarditis Lymphadenitis (occupational) Bacteremia Urinary tract infections (females) Wound infections, respiratory tract infections, foreign body infections, respiratory tract infections Catheter infections, bacteremia, endocarditis, wound infections Respiratory diphtheria, cutaneous infections Urinary tract infections, bacteremia, wound infections Bacteremia, foreign body infections, urinary tract infections Bacteremia, foreign body infections, malodorous feet Wound infections, bacteremia Cutaneous infection with erythrasma Endocarditis, bacteremia, respiratory tract infections Bacteremia, wound infections, cholecystitis Foreign body infections, bacteremia Bacteremia, foreign body infections, wound infections Pharyngitis in older children/young adults, wound and tissue infections Abscess formation (together with mixed anaerobic flora) Abscess formation, wound and soft tissue infections Bacterial vaginosis, endometritis, postpartum sepsis

480 n

BACTERIOLOGY

ethnic groups (e.g., indigenous peoples in the Americas and Australia). In the 1990s, diphtheria reemerged in the states of the former Soviet Union with a decline in the next decade. However, despite increased global travel activities, only a few imported cases have been reported by countries with well-developed health care systems. The main manifestation of diphtheria is an upper respiratory tract illness with a sore throat, dysphagia, lymphadenitis, low-grade fever, malaise, and headache. A nasopharyngeal adherent membrane which may occasionally lead to obstruction is characteristic. The severe systemic effects of diphtheria include myocarditis, neuritis, and kidney damage caused by the C. diphtheriae exotoxin, which is encoded by a bacteriophage carrying the tox gene. C. diphtheriae may also cause cutaneous diphtheria or endocarditis (with either toxin-positive or toxin-negative strains). Some people with poor hygienic standards (e.g., drug and alcohol abusers) are prone to colonization (on the skin more often than in the pharynx) or, rarely, significant infection by C. diphtheriae strains, which are nontoxigenic (50).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS In general, coryneform bacteria do not need special handling when samples are collected.

that the sensitivity of the microscopic examination is limited. Antigen assays for the direct detection of coryneform bacteria are not recommended. As described previously, C. diphtheriae, C. ulcerans, and C. pseudotuberculosis are the only species able to harbor the bacteriophage which carries the diphtheria tox gene and potentially produce diphtheria toxin. PCR-based, direct detection systems for the diphtheria tox gene have been described, using conventional methods to detect fragment A and/or the entire tox gene (51, 52) or fragment A and B subunits of the tox gene (53). The system described by Nakao et al. had the highest sensitivity when Dacron polyester-tipped swabs were used and when silica gel packages were stored at 4°C rather than at room temperature. Conventional PCR detection of the regulatory dtxR gene has been evaluated (54). Detection of the tox gene using a real-time platform has been outlined (55–57). Real-time detection using primers targeting the C. diphtheriae tox gene for C. ulcerans strains has required modifications (56–58). Direct detection of the diphtheria tox or dtxR as the sole test of clinical specimens has not been recommended, as expression of diphtheria toxin must be demonstrated, so microbiological culture is essential for confirming diphtheria (59).

C. diphtheriae

ISOLATION PROCEDURES The diagnosis of diphtheria is primarily a clinical one. The physician should notify the receiving laboratory immediately of suspected diphtheria. In case of respiratory diphtheria, material for culture should be obtained on a swab (either a cotton- or a polyester-tipped swab) from the inflamed areas in the nasopharynx. Multisite sampling (nasopharynx) is thought to increase sensitivity. If membranes are present and can be removed (swabs from beneath the membrane are most valuable), they should also be sent to the microbiology laboratory (although C. diphtheriae might not be culturable from those in every instance). Nasopharyngeal swabs should be obtained from suspected carriers. It is preferable that the swabs are immediately transferred to the microbiology laboratory for culturing. If the swabs must be sent to the laboratory, semisolid transport media (e.g., Amies) ensure the maintenance of the bacteria. All coryneform bacteria are relatively resistant to drying and moderate temperature changes. Material from patients with suspected cases of wound diphtheria can be obtained by swab or aspiration. Long-term preservation in skim milk at −70°C is applicable to all coryneform bacteria. The same skim milk tube, except for those containing lipophilic corynebacteria, can be thawed and put into the freezer again, and this can be done several times (G. Funke, unpublished data). For both nonlipophilic and lipophilic coryneforms, good results were observed with Microbank tubes (Pro Lab Diagnostics, Austin, TX) (Funke, unpublished). The advantage of using these tubes is that individual beads can be taken out of the tube. Coryneform bacteria can also be stored for decades when they are kept lyophilized in an appropriate medium (e.g., 0.9% NaCl containing 2% bovine serum albumin).

DIRECT EXAMINATION After the appropriate isolation media have been inoculated (see below under “Isolation Procedures”), the swabs taken from diphtheritic membranes may be subjected to Neisser or Loeffler methylene blue staining (positive if metachromatic granules [polar bodies] are seen). However, it is noteworthy

Coryneform bacteria including C. diphtheriae can be readily isolated from a 5% sheep blood agar (SBA)-based selective medium containing 100 μg of fosfomycin per ml (plus 12.5 μg of glucose-6-phosphate per ml), since nearly all coryneforms (except Actinomyces spp. and D. hominis) are highly resistant to this compound (60). It is also possible to put disks containing 50 μg of fosfomycin (plus 50 μg of glucose-6-phosphate [already incorporated in the disk]) (BD Diagnostics, Sparks, MD) on an SBA or a colistin-nalidixic acid (CNA) plate and then examine the colonies which grow around the disk. Selective media for coryneform bacteria containing 50 to 100 μg of furazolidone (Sigma, St. Louis, MO)/ml have also been described. If lipophilic corynebacteria like C. jeikeium or C. urealyticum are sought, then 0.1 to 1.0% Tween 80 (Merck, Darmstadt, Germany) could be added to an SBA plate (add Tween 80 before pouring the medium). Additional methods to demonstrate lipophilia are described below. Medically relevant coryneforms described to date do not grow on MacConkey agar. However, if “coryneform” bacteria are recovered from this medium, they should be examined carefully to rule out rapidly growing mycobacteria. With very few exceptions (some arthrobacters, microbacteria, and curtobacteria, which have optimal growth temperatures of between 30 and 35°C), the medically relevant coryneform bacteria all grow at 37°C. It is desirable to culture specimens for coryneform bacteria in a CO2-enriched atmosphere, since some taxa, e.g., Rothia, Arcanobacterium, and Trueperella, grow much better under those conditions. Nearly all medically relevant coryneform bacteria grow within 48 h, so primary culture plates should not be incubated longer than that. However, if liquid media are used (e.g., for specimens from normally sterile body sites), these should be checked after 5 days by Gram staining for the presence of coryneform bacteria (only if growth is observed with the naked eye) before they are discarded. It is recommended that urine specimens be incubated for longer than 24 h to check for the presence of C. urealyti-

28. Coryneform Gram-Positive Rods n 481

cum but only when patients are symptomatic or have alkaline urine or struvite crystals in their urine sediment.

C. diphtheriae The primary plating media for the cultivation of C. diphtheriae should be SBA plus one selective medium (e.g., cystine-tellurite blood agar [CTBA] or freshly prepared Tinsdale medium) (59, 61). If silica gel is used as a transport medium, the desiccated swabs need to be additionally incubated overnight in broth (supplemented with either plasma or blood), which should then be streaked onto the primary plating medium. The plates are read after 18 to 24 h of incubation at 37°C, preferably in a 5% CO2-enriched atmosphere. Tellurite inhibits the growth of many noncoryneform bacteria, but even a few C. diphtheriae strains are sensitive to potassium tellurite and will therefore not grow on CTBA but may grow on SBA. It is noteworthy that growth on CTBA and tellurite reduction are not specific for C. diphtheriae, since many other coryneforms may also produce black (albeit smaller) colonies. The best medium for direct culturing of C. diphtheriae is probably Tinsdale medium (61). However, the limitations of Tinsdale medium are its relatively short shelf life (90

a

Adapted from reference 117. In all cases, one full sweep refers to scanning the full length (2 cm) of a smear 1 cm wide by 2 cm long.

b

×450 0 1–2, 70 (1.5) 2–18, 50 (1) 4–36, 10 4–36 >36

552 n

BACTERIOLOGY

available for the recovery of mycobacteria include nonselective and selective ones (117, 130, 132), the latter containing one or more antibiotics to prevent overgrowth by contaminating bacteria or fungi. Broth media are preferred for a rapid initial isolation of mycobacteria.

Solid Media Egg-Based Media Egg-based media contain whole eggs or egg yolk, potato flour, salts, and glycerol and are solidified by inspissation. These media have a good buffer capacity, a long shelf life (several months, when refrigerated), and support good growth of most mycobacteria. Also, materials in the inoculum or medium toxic to mycobacteria are neutralized. Disadvantages of these media include variations from batch to batch depending on the quality of the eggs used, difficulties in discerning colonies from debris, and the inability to achieve accurate and consistent drug concentrations for susceptibility testing. When egg-based media become contaminated, they may liquefy. Of the egg-based media, L-J medium is most commonly used in clinical laboratories. The concentration of malachite green in standard acid-fast media (e.g., L-J) was selected to maximize the growth of mycobacteria while inhibiting other microorganisms. In general, it recovers M. tuberculosis well but is not as reliable for the recovery of other species. M. bovis, for instance, grows less well on L-J medium, but growth is stimulated if glycerol is replaced by pyruvate. In contrast to most members of the M. tuberculosis complex, M. bovis is able to grow in a reduced-O2 atmosphere. M. genavense fails to grow on L-J medium. Good recovery of M. ulcerans is obtained on L-J medium with glycerol (150, 173). Petragnani medium contains about twice as much malachite green as does L-J medium and is most commonly used for recovery of mycobacteria from heavily contaminated specimens. American Trudeau Society medium contains a lower concentration of malachite green than L-J medium and is, therefore, more easily overgrown by contaminants, but the growth of mycobacteria is less inhibited, resulting in earlier growth of larger colonies.

Agar-Based Media Compared to egg-containing media, agar-based media are chemically better defined. They do not readily support the growth of contaminants; however, the plates are expensive to prepare and their shelf life is relatively short (1 month in the refrigerator). Care should be exercised in the preparation, incubation, and storage of the media, because excessive heat or light exposure may result in deterioration and in the release of formaldehyde, which is toxic to mycobacteria. Agar-based media are transparent and provide a ready means of detecting the early growth of microscopic colonies easily distinguished from inoculum debris. Colonies may be observed in 10 to 12 days, in contrast to 18 to 24 days with egg-based media. Microscopic examination can be performed by simply turning over the plate and examining it by focusing on the agar surface through the bottom of the plate at a ×10 to ×100 magnification. This may provide both earlier detection of growth than unaided visual examination and presumptive identification of the species of mycobacteria present. The use of thinly poured 7H11 agar plates (10 by 90 mm; Remel, Lenexa, KS) facilitates this process, as microcolonies are visible after 11 days (174). Agar-based media can also be used for susceptibility testing (175). In the quest for more-rapid and affordable growth techniques, microcolony-based culture methods have been

developed. However, as shown by a recent systematic review, there is sufficient evidence neither for the feasibility and costs of implementation of these tests for growth detection and drug susceptibility testing nor for the impact on patient outcomes (176). Middlebrook medium contains 2% glycerol, which enhances the growth of MAC organisms. Nonantibiotic supplements may be helpful for the recovery of other mycobacteria and in special situations. Addition of 0.2% pyruvic acid is recommended if M. bovis is suspected, and 0.25% Lasparagine or 0.1% potassium aspartate added to 7H10 agar maximizes the production of niacin. Addition of 0.1% enzymatic hydrolysate of casein to the Middlebrook 7H11 formulation (the only difference from 7H10) improves the recovery of isoniazid-resistant strains of M. tuberculosis. M. genavense fails to grow on 7H11 agar as well. However, Middlebrook 7H11 agar supplemented with mycobactin J (Allied Monitor, Fayette, MO) supports the growth of M. genavense, as do microaerophilic conditions, the radiometric Bactec 7H12 pyrazinamide test medium (101), or addition of blood and charcoal to acidified Middlebrook agar (177).

Selective Media The addition of antimicrobial agents may be helpful in eliminating the growth of contaminating organisms. If a selective medium is used for a particular specimen, it should not be used alone but in conjunction with a nonselective agar- or egg-based medium. Egg-based selective media include L-J Gruft medium with penicillin and nalidixic acid and Mycobactosel L-J medium with cycloheximide, lincomycin, and nalidixic acid. Mitchison selective 7H11 medium and its modifications contain carbenicillin (especially useful for inhibiting pseudomonads), polymyxin B, trimethoprim lactate, and amphotericin B.

Heme-Containing Medium for the Growth of M. haemophilum M. haemophilum will grow on egg- or agar-based media only if they are supplemented with hemin, hemoglobin, or ferric ammonium citrate (105). Thus, clinical specimens should be inoculated either on chocolate agar or on media with supplements (e.g., Middlebrook 7H10 agar with hemolyzed sheep erythrocytes, hemin, or a factor X disk, or on L-J medium containing 1% ferric ammonium citrate) to enhance the recovery of this organism. Broth media should be similarly supplemented. M. haemophilum can be isolated from the MGIT (178), whereas the MB/BacT Alert 3D system is less optimal for this organism (179). As a whole, M. haemophilum infections may be underrecognized because of its predilection for a low incubation temperature (30°C) and its unique nutritional requirements.

Liquid Media Broth media may be used for both primary isolation and subculturing of mycobacteria. Cultures based on liquid media yield significantly more rapid results than solid-medium-based cultures. Also, isolation rates for mycobacteria are higher. Middlebrook 7H9 and Dubos Tween-albumin broths are commonly used for subculturing stock strains of mycobacteria and preparing the inoculum for drug susceptibility tests and other in vitro tests. 7H9 broth is used as the basal medium for several biochemical tests. Tween 80 can be added to liquid media and acts as a surfactant that allows the dispersal of clumps of mycobacteria, resulting in a more homogeneous growth. At present, the commercially available culture systems marketed for the isolation of mycobacteria range from simple

30. Mycobacterium: General Characteristics n

tubes, such as the MGIT (Becton, Dickinson Microbiology Systems), to the fully automated systems (e.g., Bactec MGIT 960 [Becton, Dickinson], ESP culture system II and VersaTREK culture system II [Trek Diagnostic Systems, Cleveland, OH], and MB/BacT Alert 3D system [bioMérieux]). These nonradiometric techniques have been tested in various studies against solid media and the radiometric Bactec 460 technique, which has long been considered the gold standard but has been discontinued by the manufacturer.

Mycobacterium Growth Indicator Tube (MGIT) The MGIT (Becton Dickinson Microbiology Systems) contains a modified Middlebrook 7H9 broth in conjunction with a fluorescence quenching-based oxygen sensor (silicon rubber impregnated with a ruthenium pentahydrate) to detect growth of mycobacteria. The large amount of oxygen initially present in the medium quenches the fluorescence of the sensor. Growth of mycobacteria or other microorganisms in the broth depletes the oxygen, and the indicator fluoresces brightly when the tubes are illuminated with UV light at 365 nm. For the manual version, a Wood’s lamp or a transilluminator can be used as the UV light source, while in the automated Bactec MGIT 960 system (see below), tubes are continuously monitored by the instrument. Prior to use, the 7H9 broth is supplemented with oleic acidalbumin-dextrose to promote the growth of mycobacteria and with polymyxin B, amphotericin B, nalidixic acid, trimethoprim, and azlocillin (PANTA) to suppress the growth of contaminants. Overall, the sensitivity and time to growth detection of the MGIT system are similar to those of the Bactec 460TB system and have been superior to solid media in clinical evaluations (180). However, contamination rates for the MGIT system are slightly higher than for the Bactec 460TB system, probably owing to the enrichments added to the MGIT broth that enhance the growth of both mycobacteria and nonmycobacterial organisms. The principal advantages of the manual MGIT system over the Bactec 460TB system include reduced opportunity for cross-contamination of cultures, no need for needle inoculation, no radioisotopes, and no need for special instrumentation other than a UV light source. Its limitations include higher contamination rates, masking of fluorescence by blood or grossly bloody specimens, and lack of compatibility with some methods of digestion and decontamination of specimens (144).

Automated, Continuously Monitoring Systems Several automated, continuously monitoring systems have been developed for the growth and detection of mycobacteria, e.g., the Bactec MGIT 960 (Becton Dickinson), the VersaTREK culture system (VersaTREK Diagnostic Systems), and the MB/BacT Alert 3D (bioMérieux). All have in common that they are no longer based on the use of radioisotopes. The MGIT system uses a fluorescence quenching-based oxygen sensor (ruthenium pentahydrate) to detect growth. The technology used in the VersaTREK culture system is based on the detection in a sealed bottle of pressure changes in the headspace above the broth medium that result from gas production or consumption due to the growth of microorganisms. The MB/BacT Alert 3D system employs a colorimetric carbon dioxide sensor in each bottle to detect the growth of mycobacteria. Each of the systems includes a broth similar to 7H9 broth supplemented with a variety of growth factors and antimicrobial agents. All three systems are FDA cleared for the isolation of mycobacteria; the VersaTREK culture system and the Bactec

553

MGIT 960 system are also cleared for drug susceptibility testing of M. tuberculosis complex organisms to various extents, though. These systems have similar levels of performance and operational characteristics. In clinical evaluations, recovery rates were similar to those of the Bactec 460TB system and superior to those of conventional solid media (Bactec MGIT 960 [181]; VersaTREK culture system [formerly the ESP culture system II] [182]; MB/BacT Alert 3D [179, 181]). In a meta-analysis of 10 published studies encompassing 1,381 strains from 14,745 clinical specimens, the Bactec MGIT 960 and Bactec 460TB systems revealed sensitivities/specificities in detecting mycobacteria of 81.5%/99.6% and 85.8%/99.9%, respectively. Combined with solid media, the sensitivities of the two systems increased to 87.7 and 89.7%, respectively (183). For some systems, time to detection of mycobacteria is similar to those of the old radiometric Bactec 460TB technique. Parrish et al. (184) demonstrated for the Bactec MGIT 960 system a shorter time to detection (13.5 versus 25.2 days) and a greater sensitivity (100% versus 66.6%) for the recovery of members of the M. tuberculosis complex than the MB BacT/Alert system. For blood specimens, the BacT/Alert MB system, the manual Myco/F Lytic medium, and the Isolator 10 lysis-centrifugation system detected M. tuberculosis in 16.4 days, 20.0 days, and 23.8 days, respectively (185). Throughout the literature, contamination rates are reported to have been higher with these new systems than with the Bactec 460TB system. However, all of them share advantages over the radiometric broth system in having no potential for cross-contamination by the instrument, being less labor-intensive, having continuous monitoring, using no radioisotopes, addressing safety more appropriately, and offering electronic data management. Since these systems monitor continuously, bottles are incubated in the instruments for their entire life in the laboratory. As a consequence, these systems are both instrument and space intensive. Some automated systems also lack the versatility of the Bactec 460TB system in that inoculation of blood is not possible, and therefore, additional instruments, e.g., the Bactec 9050 apparatus with Bactec Myco/F Lytic medium, have to be used for this purpose; alternatively, the bottles can be put into the standard Bactec 9240 or FX instrument. The same holds for the incubation of cultures harboring mycobacteria with a lower optimum temperature, such as M. chelonae, M. haemophilum, M. marinum, or M. ulcerans. Susceptibility testing applications for the primary and second-line antituberculosis drugs are available for the Bactec MGIT 960 system and VersaTREK system (formerly ESP culture system II [175]; see chapter 76). Whenever one works with automated continuously monitoring systems, mycobacterial growth can go undetected. As demonstrated by Pena et al. (186), 1,323 (13%) of 10,263 tested specimens were culture positive for mycobacteria, but approximtely 1% of instrument-negative MGIT cultures contained mycobacterial growth when tubes had been inspected visually for clumps before the tubes were discarded after 8 weeks.

Medium Selection Medium selection for the isolation of mycobacteria and culture reading schedules are usually based on personal preferences and/or laboratory tradition. Both should be optimized for the most rapid detection of positive cultures and identification of mycobacterial isolates. The variety of media and methods available today is sufficient to permit laboratories to develop an algorithm that is optimal for their patient

554 n

BACTERIOLOGY

population and administrative needs. Workload, financial resources, and, in particular, the limited amounts of processed sediments are, however, restraining factors in working with too many different types of media. Thus, cultivation of mycobacteria always involves a compromise. Today, it is generally accepted that the use of a liquid medium in combination with at least one solid medium is essential for good laboratory practice in the isolation of mycobacteria (130). Addition of a solid medium is advantageous for those strains which occasionally do not grow in liquid medium, aids in the detection of mixed mycobacterial infections, and can serve as a backup for broth, which has a higher contamination rate. All positive cultures, even if identified directly from the broth, must be subcultured to solid media to detect mixed cultures and to correlate direct identification results with colony morphology. The nonradiometric growth systems cannot serve as stand-alone culture systems for mycobacteria for the reasons stated above. Detection of colonies on solid medium certainly offers several advantages over detection of growth in broth, because colonial morphology can provide clues to identification and facilitate the selection of confirmatory tests. However, smears from broth-based systems can sometimes provide microscopic clues, such as cord formation (see above), although the reliability for presumptive identification of M. tuberculosis should be applied with caution since the phenomenon is also observed with some NTM species (158, 159, 160).

Incubation Temperature The optimum incubation temperature for most cultures is 35 to 37°C. Exceptions to this include cultures obtained from skin and soft tissue suspected of containing M. marinum, M. ulcerans, M. chelonae, or M. haemophilum, which have lower optimum temperatures. For such specimens, a second set of media has to be inoculated and incubated at 25 to 33°C. Lower temperatures increase detection time. The newer automated liquid-medium-based culture systems do not offer the possibility of incubating at temperatures lower than 36 ± 1°C.

Atmosphere Five percent to 10% CO2 in air stimulates the growth of mycobacteria in primary isolation cultures using conventional media. Middlebrook agar requires a CO2 atmosphere to ensure growth, while it is necessary to incubate egg media under CO2 for only the first 7 to 10 days after inoculation, i.e., during the log phase of growth. Subsequently, L-J cultures can be removed to ambient-air incubators if space is limited. In the absence of CO2 incubators, plates may be incubated in commercially available bags with CO2-generating tablets. Candle extinction jars are unacceptable for use in mycobacteriology laboratories because the oxygen tension is less than that required for the growth of mycobacteria. Broth systems usually do not require incubation at increased CO2 concentrations.

suspicion of tuberculosis. Plates should be incubated with the medium side down until the entire inoculum has been absorbed. Once this has happened, media should be incubated inverted in CO 2-permeable polyethylene bags or sealed with CO2-permeable shrink-seal bands to prevent them from drying up during the incubation period. Tubed media should be incubated in a slanted position with the screw caps loose for at least a week until the inoculum has been absorbed; they can then be incubated upright if space is at a premium. Caps on the tubes should be tightened at 2 to 3 weeks to prevent desiccation of the media. Specimens from skin lesions should be incubated for 8 to 12 weeks if M. ulcerans is suspected.

Reading Schedule Since many mycobacteria are slowly growing organisms, cultures can be examined less frequently than routine bacteriologic cultures. All solid media should be examined within 3 to 5 days after inoculation to permit early detection of rapidly growing mycobacteria and to enable prompt removal of contaminated cultures. Young cultures (up to 4 weeks of age) should be examined twice a week, whereas older cultures could be examined at weekly intervals. Use of a hand lens for opaque media and a microscope for agar media will facilitate early detection of microcolonies. The manual MGIT may be inspected for growth daily for the first 1 to 2 weeks. Afterwards, it should be inspected twice weekly or weekly for growth. When using one of the nonradiometric continuously monitoring systems, laboratory personnel are automatically alerted by the instrument if a specimen turns positive. Irrespective of the system used, the acid fastness of the organism has to be confirmed by smear staining. Also, it is highly advisable to subculture the broth on a sheep blood or chocolate agar plate to rule out contaminants. Once growth of AFB is detected, susceptibility testing can be performed, always according to the instructions specified by the manufacturers.

Reporting Traditionally, solid culture media are being kept for up to 8 weeks. Even though liquid media detect mycobacteria much earlier, most incubation protocols still require a minimum of 6 weeks (130). A large multicenter evaluation involving >1,500 positive cultures (among them, 466 M. tuberculosis isolates) stressed that laboratories using MGIT may issue reports of no growth of M. tuberculosis as early as at 4 weeks (187). This was confirmed by Pfyffer and Wittwer (188); 58.3% of all mycobacteria were detected within 14 days, 37.5% were detected within 21 days, and 4.2% were detected within 28 days if a combination of solid media (L-J/Middlebrook) and liquid medium (Bactec MGIT 960) were utilized. In this context, 50% of species in the M. tuberculosis complex were detected within 14 days, and 50% were detected within 21 days. As much as 70% of NTM appeared within 14 days, 20% were detected within 21 days, and 10% were detected within 28 days.

Storage of Positive Cultures Time Mycobacterial cultures on solid and in liquid media are generally held for 6 to 8 weeks before being discarded as negative. Specimens with positive smears that are culture negative should be held for an additional 4 weeks. The same should be done for culture-negative specimens that were positive for mycobacteria by one of the nucleic acid-based amplification assays or for cases for which there is a persisting

Positive cultures may be kept at room temperature for several weeks. If subcultured, they may be saved at room temperature for several months. Solid cultures have to be sealed to avoid dehydration of the medium. The CLSI (130) recommends that cultures that may be needed for possible followup in the future be frozen at –70°C (a minimum of 1 year is recommended) or cryopreserved as suggested by Shu et al. (189).

30. Mycobacterium: General Characteristics n

IMMUNODIAGNOSTIC TESTS FOR TUBERCULOSIS Historically, the first immunodiagnostic test was the TST. The shortcomings of this test are well known and include the inability to distinguish active tuberculosis disease from past sensitization by BCG, unknown predictive values, and cross-reaction with NTM. Over time, much effort has been devoted to the development of serological tests for the diagnosis of tuberculosis, but no test has found widespread clinical use. The sensitivity and specificity of serological tests with crude antigen preparations are too low for clinical application. In contrast, the two commercially available whole-blood IGRAs have become a more promising, though not perfect, tool to detect tuberculosis infection, in particular, if TST remains equivocal. Both tests are FDA approved but not for testing individuals under the age of 17, individuals with immunedepression due to AIDS, persons taking immunosuppressive drugs (e.g., anti-tumor necrosis factor or corticosteroids), or transplant recipients. Two test systems, the T-Spot.TB (Oxford Immunotec, Oxford/United Kingdom) and the QuantiFERON gold in-tube (QFNG-IT; Cellestis, Victoria/ Australia) tests, are not affected by BCG vaccination, do not cross-react with the majority of NTM, and are less prone to variability and subjectivity associated with placing and reading of the TST. Also, individuals to be tested have to see a doctor or health care personnel only once. As shown by a recent systematic review of studies performed in high-, middle-, and low-income countries, the higher per-test cost of IGRAs may be compensated for by low postscreening costs (medical attention, chest X rays, chemoprevention), given the higher specificity of the IGRAs than that of conventional TST (190). In particular, there was strong evidence of the cost-effectiveness of the IGRAs in screening risk groups, such as immigrants from high-incidence countries, close contacts, and health care workers. Disagreement between QFNG-IT and TST results as well as the high reversion rate with QFNG-IT, however, raised concerns about the effectiveness of QFNG-IT as a sole screening test for health care workers (191). Although both tests measure T cell IFN-γ responses to two or three M. tuberculosis-specific antigens (ESAT-6, CFP-10, TB 7.7) over a 16- to 24-h incubation period, they are based on different technologies. The T-SPOT.TB assay is based on the enzyme-linked immunosorbent spot (ELISPOT) methodology and requires the isolation and incubation of peripheral blood mononuclear cells (PBMC) and the standardization of 250,000 PBMC in each of its test wells. The assay requires, overall, two working days and may be more laborious than the QFNG-IT. Nevertheless, the use of a standardized number of washed PBMC may represent another advantage. In contrast, the QFNG-IT assay has technical advantages over the T-SPOT.TB assay, since the stimulation of a T cell IFN-γ response in whole blood is performed in tubes precoated with the M. tuberculosis antigens. Also, the enzyme-linked immunosorbent assay (ELISA) is simple to perform and requires one working day. Since background noise may occur, a “nil” control is required to adjust for this background, as well as for heterophile antibody effects and nonspecific IFN-γ in blood samples. The reproducibility of QFNG-IT assay results in duplicate tests is excellent (192). It is important to stress that neither of these new tests distinguishes between latent and active infection. To date, there are several guidelines available, among them the one for the United States (193). A large number of publications

555

focus on the performance characteristics of each test compared to TST. As a whole, IGRAs are better correlated with the intensity of tuberculosis exposure than the TST (194). There are several systematic reviews and meta-analyses which document the performance characteristics of the two types of IGRAs in both immunocompetent and immunocompromized patients. In a head-to-head comparison of the QFNG-IT and the T-SPOT.TB assays, Pai et al. (195) concluded that the QFNG-IT test has a specificity of 99% (T-SPOT.TB test, 96%) among non-BCG-vaccinated participants and a specificity of 96% (T-SPOT.TB test, 93%) among BCG-vaccinated participants, while Higuchi et al. (196) reported equal specificities for the two assays. For latent tuberculosis, specificity varied between 98 and 100%. In immunocompetent adults, the negative predictive values for progression to tuberculosis within 2 years were 97.8% for the T-SPOT.TB assay and 99.8% for the QFNG-IT assay (197). In contrast, it is well understood that IGRAs have limited accuracy in diagnosing active tuberculosis, as evidenced by a meta-analysis of 884 studies (198). This also holds true for HIV-positive individuals (199). Also, QFNGIT results do not offer much value for treatment monitoring of tuberculosis disease (200). Diel et al. (201) compared both IGRAs in TST-positive persons recently exposed to pulmonary tuberculosis cases. In that study, factors independently influencing the risk of M. tuberculosis infection and their interactions with each other were evaluated by multivariate analysis. There were five variables which significantly predicted a positive IGRA result, i.e., age, AFB positivity of the source case, cough, cumulative exposure time, and foreign origin of the patient. There was excellent agreement between the two assays (93.9%, kappa = 0.85), with QFNG-IT finding 30.2% of contacts positive and T-SPOT.TB finding 28.7% of them. Again, the IGRAs were more accurate indicators of the presence of latent tuberculosis than the TST (201). In HIV-positive asymptomatic individuals (n = 286), both the QFNG-IT and T-SPOT.TB assays were more sensitive than the TST (20.0% and 25.2%, respectively, compared with 12.8% for the TST) but seemed, as a whole, to be less sensitive than in immunocompetent patients (202). The value of TST and serial QFNG-IT tests in patients with rheumatic diseases during long-term treatment with tumor necrosis factor blockers has not been defined to date (203). The performance of IGRAs in children is less understood. Without having the inconveniences and complications associated with the TST, IGRAs are acceptable substitutes for it. The sensitivity and specificity of IGRAs are, however, not significantly higher than the values observed for the TST (204). In children with latent tuberculosis, the agreement between the QFNG-IT and T-SPOT.TB assays was very good (92%), with moderate agreement between the TST and the QFNG-IT assay (77%) and the TST and the T-SPOT.TB assay (75%) (205). Available data suggest that the TST and IGRAs have similar levels of accuracy for the detection of tuberculosis infection or the diagnosis of disease in children (206). Since the experience with IGRAs is still limited, longitudinal studies are needed to define their predictive values, especially in children and high-risk populations (207). A large meta-analysis has pointed out very clearly present-day limitations of IGRAs, such as with their performance in children, in immunocompromized persons, and in the elderly (208). Other problems concern altered performance characteristics of the assays in conjunction with ethnicity

556 n BACTERIOLOGY

(209), specifically with the phenomena of conversions, reversions, and nonspecific variations in serial testing (210– 212). Assessment of within-subject IGRA variability is important in establishing thresholds for conversions and reversions. With the QFNG-IT assay, Detjen et al. (192) observed considerable intraindividual variability in serial analyses within 3 days. van Zyl-Smit et al. (213) concluded from their study that a 3-spot or 80% IFN-γ response variation on either side of the baseline values explains 95% of the short-term variability and may be useful for interpreting conversions, reversions, and values close to the cutoff point. Therefore, these authors have proposed a borderline or uncertainty zone for both tests (4 to 8 spots [inclusive] for the T-SPOT.TB assay and 0.2 to 0.7 IU/ml for the QFNGIT assay) instead of strictly adhering to the manufacturerdefined cutoff value (T-SPOT.TB assay, > 6 spots; QFNGIT test, >0.35 IU/ml). Basically, the problem of indeterminate results occurs with both IFN-γ release assays. In HIV-infected individuals, the T-SPOT.TB assay provided more indeterminate results than the QFNG-IT test (8 versus 1/256, P < 0.01) (202), similar to what has been confirmed by others (14% versus 1.8%) (214). If preanalytical errors can be excluded, indeterminate results appear to be dependent on the number of CD4 cells, inasmuch as patients with a CD4 count of ≤200 cells/ml were significantly more likely to have an indeterminate result (202, 214). In children 24 h). Mycobacteria remain viable for 8 days in the solution.

Reagents CPC digestant-decontaminant. Dissolve 10 g of CPC and 20 g of NaCl in 1,000 ml of distilled water. The solution is self-sterilizing and remains stable if protected from light, extreme heat, and evaporation. Dissolve with gentle heat any crystals that might form in the working solution. Other reagents used in processing include sterile water and sterile saline or 0.2% sterile BSA fraction V.

Procedure 1. Collect 10 ml or less of sputum in a 50-ml screw-cap centrifuge tube. 2. Inside a BSC, add an equal volume of CPC-NaCl, cap securely, and shake by hand until the specimen liquefies. 3. Package the specimen appropriately as specified by current postal regulations, and send it to a processing laboratory. 4. Upon receipt in the processing laboratory (allow at least 24 h for digestion/decontamination to be completed), dilute the digested/decontaminated specimen to the 50-ml mark with sterile distilled water and recap securely. Invert the tube several times to mix the contents. 5. Centrifuge at ≥3,000 × g for 15 min, decant the supernatant fluid, and suspend the sediment in 1 to 2 ml of sterile water, saline, or 0.2% BSA fraction V.

560 n

BACTERIOLOGY

6. Inoculate the resuspended sediment onto egg medium, and make a smear.

12.

Sulfuric Acid Method Principle The sulfuric acid method may be useful for urine and other body fluids that yield contaminated cultures when processed by one of the alkaline digestants.

13.

Reagents 4% sulfuric acid 4% sodium hydroxide Sterile distilled water Phenol red indicator

14.

15.

Procedure 1. Centrifuge the entire specimen for 30 min at ≥3,000 × g. This may require several tubes. 2. Decant the supernatant fluids; pool the sediments if several tubes were used for a single specimen. 3. Add an equal volume of 4% sulfuric acid to the sediment. 4. Vortex and let stand for 15 min at room temperature. 5. Fill the tube to the 50-ml mark with sterile water. 6. Centrifuge at ≥3,000 × g for 15 min and decant the supernatant. 7. Add 1 drop of the phenol red indicator and neutralize with 4% NaOH until a persistent pale pink color forms. 8. Inoculate the media and make a smear.

16. 17.

18.

REFERENCES 1. World Health Organization. 2012. Global tuberculosis report. WHO Report 2012. World Health Organization, Geneva, Switzerland. http://www.who/int/tb/publications/ global_report/en/. 2. American Thoracic Society. 2007. Diagnosis, treatment, and prevention of nontuberculous mycobacterial diseases. Am J Respir Crit Care Med 175:367–416. 2. Falkinham JO. 1996. Epidemiology of infection by nontuberculous mycobacteria. Clin Microbiol Rev 9:178–215. 4. Falkinham JO. 2013. Ecology of nontuberculous mycobacteria—where do human infections come from? Semin Respir Crit Care Med 34:95–102. 5. Tortoli E. 2003. Impact of genotypic studies in mycobacterial taxonomy: the new mycobacteria of the 1990s. Clin Microbiol Rev 16:319–354. 6. Primm TP, Lucero CA, Falkinham JO, III. 2004. Health impacts of environmental mycobacteria. Clin Microbiol Rev 17:98–106. 7. Wayne LG, Kubica GP. 1986. Mycobacteria, p 1435– 1457. In Sneath PHA (ed), Bergey’s Manual of Systemic Bacteriology, vol 2. William’s & Wilkins, Baltimore, MD. 8. Conville PS, Witebsky FG. 2005. Nocardia and other aerobic actinomycetes, p 1137–1180. In Boriello SP, Murray PR, Funke G (ed), Topley & Wilson’s Microbiology and Microbial Infections, Bacteriology, 10th ed, vol 2. Hodder Arnold, London, United Kingdom. 9. Butler RW, Floyd MM, Brown JM, Toney SR, Daneshvar MI, Cooksey RC, Carr J, Steigerwalt AG, Charles N. 2005. Novel mycolic acid-containing bacteria in the family Segniliparaceae fam. nov., including the genus Segniliparus gen. nov., with descriptions of Segniliparus rotundus sp. nov. and Segniliparus rugosus sp. nov. Int J Syst Evol Microbiol 55:1615–1624. 10. Ghosh J, Larsson P, Singh B, Pettersson BM, Islam NM, Sarkar SN, Dasgupta S, Kirsebom LA. 2009. Sporulation in mycobacteria. Proc Natl Acad Sci U S A 106:10781– 10786. 11. Kremer L, Besra GS. 2005. A waxy tale, by Mycobacterium tuberculosis, p 287–305. In Cole ST, Eisenach KD, McMur-

19.

20.

21. 22.

23.

24.

25.

26.

ray DN, Jacobs WR, Jr (ed), Tuberculosis and the Tubercle Bacillus. ASM Press, Washington, DC. Wheeler PR, Blanchard JS. 2005. General metabolism and biochemical pathways of tubercle bacilli, p 309–339. In Cole ST, Eisenach KD, McMurray DN, Jacobs WR, Jr (ed), Tuberculosis and the Tubercle Bacillus. ASM Press, Washington, DC. Mitscherlich E, Marth EH. 1984. Microbial Survival in the Environment—Bacteria and Rickettsiae Important in Human and Animal Health, p 232–266. Springer-Verlag, Berlin, Germany. World Health Organization. 2012. Tuberculosis Laboratory Biosafety Manual. WHO, Geneva, Switzerland. http://apps .who.int/iris/bitstream/10665/77949/1/9789241504638_ eng.pdf. Chosewood LC, Wilson DE (ed). 2007. Biosafety in Microbiological and Biomedical Laboratories, 5th ed. Centers for Disease Control and Prevention, Washington, DC. http:// www.cdc.gov/od/ohs. Fleming DO, Hunt DL. 2006. Biological Safety: Principles and Practices, 4th ed. ASM Press, Washington, DC. Allen S, Batungwanayo J, Kerlikowske K, Lifson AR, Wolf W, Granich R, Taelman H, van de Perre P, Serufilira A, Bogaerts J. 1992. Two-year incidence of tuberculosis in cohorts of HIV-infected and uninfected urban Rwandan women. Am Respir Dis 146:1439–1444. Brown-Elliott BA, Wallace RJ, Tichindelean C, Sarria JC, McNulty S, Vasireddy R, Bridge L, Mayall CG, Turenne C, Loeffelholz M. 2011. Five-year outbreak of community- and hospital-acquired Mycobacterium porcinum infections related to public water supplies. J Clin Microbiol 49:4231–4238. Wallace R, Iakhiaeva E, Williams MD, Brown-Elliott BA, Vasireddy R, Lande L, Peterson DD, Sawicki J, Kwait R, Tichenor WS, Turenne C, Falkinham JO. 2013. Absence of Mycobacterium intracellulare and the presence of Mycobacterium chimaera in household water and biofilm samples of patients in the United States with Mycobacterium avium complex respiratory disease. J Clin Microbiol 51:1747–1752. Marshall HM, Carter R, Torbey MJ, Minion S, Tolson C, Sidjabat HE, Huygens F, Hargreaves M, Thomson RM. 2011. Mycobacterium lentiflavum in drinking water supplies, Australia. Emerg Infect Dis 17:395–402. Hernandez-Garduno E, Elwood K. 2012. Nontuberculous mycobacteria in tap water. Emerg Infect Dis 18:353. Brosch R, Gordon SV, Marmiesse M, Brodin P, Buchrieser C, Eiglmeier K, Garnier T, Gutierrez C, Hewinson G, Kremer K, Parsons LM, Pym AS, Samper S, van Soolingen D, Cole ST. 2002. A new evolutionary scenario for the Mycobacterium tuberculosis complex. Proc Natl Acad Sci U S A 99:3684–3689. Gagneux S, DeRiemer K, Van T, Kato-Maeda M, de Jong BC, Narayanan S, Nicol M, Niemann S, Kremer K, Gutierrez MC, Hilty M, Hopewell PC, Small PM. 2006. Variable host-pathogen compatibility in Mycobacterium tuberculosis. Proc Natl Acad Sci U S A 103:2869– 2873. Gutierrez MC, Brisse S, Brosch R, Fabre M, Omaïs B, Marmiesse M, Supply P, Vincent V. 2005. Ancient origin and gene mosaicism of the progenitor of Mycobacterium tuberculosis. PLoS Pathog 1(1):e5. Gandhi NR, Shah NS, Andrews JR, Vella V, Moll AP, Scott M, Weissman D, Marra C, Lalloo UG, Friedland GH. 2010. HIV coinfection in multidrug- and extensively drug-resistant tuberculosis results in high early mortality. Am J Respir Crit Care Med 181:80–86. Hopewell PC, Jasmer RM. 2005. Overview of clinical tuberculosis, p 15–31. In Cole ST, Eisenach KD, McMurray DN, Jacobs WR, Jr (ed), Tuberculosis and the Tubercle Bacillus. ASM Press, Washington, DC.

30. Mycobacterium: General Characteristics n 27. Cole ST, Brosch R, Parkhill J, Garnier T, Churcher C, Harris D, Gordon SV, Eiglmeier K, Gas S, III, Barry CE, Tekaia F, Badcock K, Basham D, Brown D, Chillingworth T, Connor R, Davies R, Devlin K, Feltwell T, Gentles S, Hamlin N, Holroyd S, Hornsby T, Jagels K, Krogh A, Maclean J, Moule S, Murphy L, Oliver K, Osborne J, Quail MA, Rajandream MA, Rogers J, Rutter S, Seeger K, Skelton J, Squares R, Squares S, Sulston JE, Taylor K, Whitehead S, Barrell BG. 1998. Deciphering the biology of Mycobacterium tuberculosis from the complete genome sequence. Nature 393:537–544. 28. Karboul A, Gey van Pittius NC, Namouchi A, Vincent V, Sola C, Rastogi N, Suffys P, Fabre M, Cataldi A, Huard RC, Kurepina N, Kreiswirth B, Ho JL, Gutierrez MC, Mardassi H. 2006. Insights into the evolutionary history of tubercle bacilli as disclosed by genetic rearrangements within a PE_PGRS duplicated gene pair. BMC Evol Biol 6:107–125. 29. Blouin Y, Hauck Y, Soler C, Fabre M, Vong R, Dehan C, Cazajous G, Massoure P-L, Kraemer P, Jenkins A, Garnotel E, Pourcel C, Vergnaud G. 2012. Significance of the identification in the Horn of Africa of an exceptionally deep branching Mycobacterium tuberculosis clade. PLoS One 7:e52841. doi:10.1371/journal.pone.0052841 30. Majoor CJ, Magis-Escurra C, van Ingen J, Boeree MJ, van Soolingen D. 2011. Epidemiology of Mycobacterium bovis disease in humans, the Netherlands, 1993–2007. Emerg Infect Dis 17:457–463. 31. Garnier T, Eiglmeier K, Camus J-C, Medina N, Mansoor H, Pryor M, Duthoy S, Grondlin S, Lacroix C, Monsempe C, Simon S, Harris B, Atkin R, Doggett J, Mayes R, Keating L, Wheeler PR, Parkhill J, Barrell BG, Cole ST, Gordon SV, Hewinson RG. 2003. The complete genome sequence of Mycobacterium bovis. Proc Natl Acad Sci U S A 100:7877–7882. 32. Behr MA, Small PM. 1999. A historical and molecular phylogeny of BCG strains. Vaccine 17:915–922. 33. Supply P, Mazars E, Lesjean S, Vincent V, Gicquel B, Locht C. 2000. Variable human minisatellite-like regions in the Mycobacterium tuberculosis genome. Mol Microbiol 36:762–771. 34. Siatelis A, Houhoula D, Papaparaskevas J, Delakas D, Tsakiris A. 2011. Detection of Bacillus Galmette-Guérin (Mycobacterium bovis BCG) DNA in urine and blood specimens after intravesical immunotherapy for bladder carcinoma. J Clin Microbiol 49:1206–1208. 35. Kanamori H, Isogami K, Hatakeyama T, Saito H, Shimada K, Uchiyama B, Aso N, Kaku M. 2012. Chest wall abscess due to Mycobacterium bovis BCG after intravesical BCG therapy. J Clin Microbiol 50:533–535. 36. Brosch R, Gordon SV, Garnier T, Eiglmeier K, Frigui W, Valenti P, Dos Santos S, Duthoy S, Lacroix S, GarciaPelayo C, Inwald JK, Golby P, Garcia JN, Hewinson RG, Behr MA, Quai MA, Churcher C, Barrell BG, Parkhill J, Cole ST. 2007. Genome plasticity of BCG and impact on vaccine efficacy. Proc Natl Acad Sci U S A 104:5596– 5601. 37. Joshi D, Harris NB, Waters R, Thacker T, Mathema B, Kreiswirth B, Sreevatsan S. 2012. Single nucleotide polymorphisms in the Mycobacterium bovis genome resolve phylogenetic relationships. J Clin Microbiol 50:3853–3861. 38. de Jong BC, Antonio M, Gagneux S. 2010. Mycobacterium africanum—review of an important cause of human tuberculosis in West Africa. PLoS Negl Trop Dis 4:e744. 39. Desmond E, Ahmed AT, Probert WS, Ely J, Jang Y, Sanders SA, Lin S-Y, Flood J. 2004. Mycobacterium africanum cases, California. Emerg Infect Dis 10:921–923. 40. Isea-Pena MC, Brezmes-Valdivieso MF, Gonzalez-Velasco MC, Lezcano-Carrera MA, Lopez-Urrutia-Lorente L, Martin-Casabona N, Monforte-Cirac ML, Palacios JJ,

41.

42.

43.

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

561

Penedo-Pallares A, Ramirez-Rosales A, Sanchez-Silos R, Tortola-Fernandez T, Vinuelas-Bayron J, Vitoria-Agreda A. 2012. Mycobacterium africanum, an emerging disease in high-income countries? Int J Tuberc Lung Dis 16:1400– 1404. Bentley SD, Comas I, Bryant JM, Walker D, Smith NH, Harris SR, Thurston S, Gagneux S, Wood J, Antonio M, Quall MA, Gehre F, Adegbola RA, Parkhill J, de Jong BC. 2012. The genome of Mycobacterium africanum West African 2 reveals a lineage-specific locus and genome erosion common to the M. tuberculosis complex. PLoS Negl Trop Dis 6:e1552. doi:10.1371/journal.pntd.0001552. Aranaz A, Liebana E, Gomez-Mampaso E, Galan JC, Cousins D, Blazquez J, Baquero F, Mateos A, Suarez G, Dominguez L. 1999. Mycobacterium tuberculosis subsp. caprae subsp. nov.: a taxonomic study of the Mycobacterium tuberculosis complex isolated from goats in Spain. Int J Syst Bacteriol 49:1263–1273. Kubica T, Rüsch-Gerdes S, Niemann S. 2003. Mycobacterium bovis subsp. caprae caused one-third of human M. bovisassociated tuberculosis cases reported in Germany between 1999 and 2001. J Clin Microbiol 41:3070–3077. Rodriguez S, Bezos J, Romero B, de Juan L, Alvarez J, Castellanos E, Moya N, Lozano F, Tariq Javed M, SaezLlorente JL, Liebana E, Mateos A, Dominguez L, Aranaz A. 2011. Mycobacterium caprae infection in livestock and wildlife, Spain. Emerg Infect Dis 17:532–535. Prodinger WM, Brandstätter A, Naumann L, Pacciarini M, Kubica T, Boschiroli LM, Aranaz A, Nagy G, Cvetnic Z, Ocepek M, Skrypnyk A, Erler W, Niemann S, Pavlik I, Moser I. 2005. Characterization of Mycobacterium caprae isolates from Europe by mycobacterial interspersed repetitive unit genotyping. J Clin Microbiol 43:4984–4992. Domogalla J, Prodinger WM, Blum H, Krebs S, Gellert S, Müller M, Neuendorf E, Sedlmaier F, Büttner M. 2013. Region of difference four (RD4) in alpine Mycobacterium caprae isolates indicates three variants. J Clin Microbiol 51:1381–1388. Oevermann A, Pfyffer GE, Zanolari P, Meylan M, Robert N. 2004. Generalized tuberculosis in Llamas (Lama glama) due to Mycobacterium microti. J Clin Microbiol 42:1818– 1821. Alvarez J, Bezos J, de Juan L, Vordermeier M, Rodriguez S, Fernandez-de-Mera IG, Mateos A, Dominguez L. 2011. Diagnosis of tuberculosis in camelids: old problems, current solutions and future challenges. Transbound Emerg Dis 59:1–10. Rüfenacht S, Bögli-Stuber K, Bodmer T, Jaunin VF, Jmaa DC, Gunn-Moore DA. 2011. Mycobacterium microti infection in the cat: a case report, literature review and recent clinical experience. J Feline Med Surg 13:195–204. Bennett AD, Lalor S, Schwarz T, Gunn-Moore DA. 2011. Radiographic findings in cats with mycobacterial infections. J Feline Med Surg 13:718–724. Palgrave CJ, Benato L, Eatwell K, Laurenson IF, Smith NH. 2012. Mycobacterium microti infection in two meerkats (Suricata suricatta). J Comp Pathol 146:278–282. van Soolingen D, van der Zanden AGM, de Haas PEW, Noordhoek GT, Kiers A, Foudraine NA, Portaels F, Kolk AHJ, Kremer K, van Embden JDA. 1998. Diagnosis of Mycobacterium microti infections among humans by using novel genetic markers. J Clin Microbiol 36:1840–1845. de Jong E, Rentenaar RJ, van Pelt R, de Lange W, Schreurs W, van Soolingen D, Sturm PDJ. 2009. Two cases of Mycobacterium microti-induced culture-negative tuberculosis. J Clin Microbiol 47:3038–3040. Panteix G, Gutierrez MC, Boschiroli ML, Rouviere M, Plaidy A, Pressac D, Porcheret H, Chyderiotis G, Ponsada M, van Oortegem K, Salloum S, Cabuzel S, Banuls AL, van de Perre P, Godreuil S. 2010. Pulmonary tubercu-

562 n

55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

65.

66.

BACTERIOLOGY

losis due to Mycobacterium microti: a study of six recent cases in France. J Med Microbiol 59:984–989. Frota CC, Hunt DM, Buxton RS, Rickman L, Hinds J, Kremer K, van Soolingen D, Colston MJ. 2004. Genome structure in the vole bacillus, Mycobacterium microti, a member of the Mycobacterium tuberculosis complex with a low virulence for humans. Microbiology 15:1519–1527. van Soolingen D, Hoogenboezem T, de Haas PEW, Hermans PWM, Koedam MA, Teppema KS, Brennan PJ, Besra GS, Portaels F, Top J, Schouls LM, van Embden JDA. 1997. A novel pathogenic taxon of the Mycobacterium tuberculosis complex, Canettii: characterization of an exceptional isolate from Africa. Int J Syst Bacteriol 47:1236–1245. Pfyffer GE, Auckenthaler R, van Embden JDA, van Soolingen D. 1998. Mycobacterium canettii, the smooth variant of M. tuberculosis, isolated from a Swiss patient exposed in Africa. Emerg Infect Dis 4:631–634. Fabre M, Koeck J-L, Le Flèche P, Simon F, Hervé V, Vergnaud G, Pourcel C. 2004. High genetic diversity revealed by variable-number tandem repeat genotyping and analysis of hsp65 gene polymorphism in a large collection of “Mycobacterium canettii” strains indicates that the M. tuberculosis complex is a recently emerged clone of “M. canettii.” J Clin Microbiol 42:3248–3255. Miltgen J, Morillon M, Koeck JL, Varnerot A, Briant JF, Nguyen G, Verrot D, Bonnet D, Vincent V. 2002. Two cases of pulmonary tuberculosis caused by Mycobacterium tuberculosis subsp. canettii. Emerg Infect Dis 8:1350– 1352. Somoskövi A, Dormandy J, Mayrer AR, Carter M, Hooper N, Salfinger M. 2009. “Mycobacterium canettii” isolated from a human immunodeficiency virus-positive patient: first case recognized in the United States. J Clin Microbiol 47:255–257. Marmiesse M, Brodin P, Buchrieser C, Gutierrez C, Simoes N, Vincent V, Glaser P, Cole ST, Brosch R. 2004. Macro-array and bioinformatics analyses reveal mycobacetrial ‘core’ genes, variation in the ESAT-6 gene family and new phylogenetic markers for the Mycobacterium tuberculosis complex. Microbiology 150:483–496. Supply P, Marceau M, Mangenot S, Roche D, Rouanet C, Khanna V, Majlessi L, Criscuolo A, Tap J, Pawlik A, Fiette L, Orgeur M, Fabre M, Parmentier C, Frigui W, Simeone R, Boritsch EC, Debrie A-S, Willery E, Walker D, Quail MA, Ma L, Bouchier C, Salvignol G, Sayes F, Cascioferro A, Seemann T, Barbe V, Locht C, Gutierrez M-C, Leclerc C, Bentley SD, Stinear TP, Brisse S, Médigue C, Parkhill J, Cruveiller S, Brosch R. 2013. Genomic analysis of smooth tubercle bacilli provides insights into ancestry and pathoadaptation of Mycobacterium tuberculosis. Nat Genet 2:172–181. Cousins DV, Bastida R, Cataldi A, Quse V, Redrobe S, Dow S, Duignan P, Murray A, Dupont C, Ahmed N, Collins DM, Butler WR, Dawson D, Rodriguez P, Loureiro J, Romano MI, Alito A, Zumarraga M, Bernardelli A. 2003. Tuberculosis in seals caused by a novel member of the Mycobacterium tuberculosis complex: Mycobacterium pinnipedii sp. nov?. Int J Syst Evol Microbiol 53:1305–1314. Jurczynski K, Lyashchenko KP, Gomis D, Moser I, Greenwald R, Moisson P. 2011. Pinniped tuberculosis in Malayan tapirs (Tapirus indicus) and its transmission to other terrestrian mammals. J Zoo Wildl Med 42:222–227. Kiers A, Klarenbeek A, Mendelts B, van Soolingen D, Koëter G. 2008. Transmission of Mycobacterium pinnipedii to humans in a zoo with marine mammals. Int J Tuberc Lung Dis 12:1469–1473. Alexander KA, Laver PN, Michel AL, Williams M, van Helden PD, Warren RM, Gey van Pittius NC. 2010. Novel Mycobacterium tuberculosis complex pathogen, M. mungi. Emerg Infect Dis 16:1296–1299.

67. Gey van Pittius NC, van Helden PD, Warren RM. 2012. Characterization of Mycobacterium orygis. Emerg Infect Dis 18:1708–1709. 68. Gey van Pittius NC, Perrett KD, Michel AL, Keet DF, Hlokwe T, Streicher EM, Warren RM, van Helden PD. 2012. Infection of African buffalo (Syncerus caffer) by oryx bacillus, a rare member of the antelope clade of Mycobacterium tuberculosis. J Wildl Dis 48:849–857. 69. van Ingen J, Rahim Z, Mulder A, Boeree MJ, Simeone R, Brosch R, van Soolingen D. 2012. Characterization of Mycobacterium orygis as M. tuberculosis complex subspecies. Emerg Infect Dis 18:653–656. 70. World Health Organization. 2012. Global leprosy situation, 2012. Wkly Epidemiol Rep 87:317–328. http:// www.who.int./wer. 71. Loughry WJ, Trumman RW, McDonough CM, Tilak MK, Garnier S, Delsuc F. 2009. Is leprosy spreading among nine-banded armadillos in the southeastern United States? J Wildl Dis 45:144–152. 72. Fite GL, Cambre FJ, Hunt MH. 1947. Procedures for demonstrating lepra bacilli in paraffin sections. Arch Pathol 43:624–625. 73. Sakamuri RM, Kimura M, Li W, Kim HC, Lee H, Madanahally K, Blackiv WC, IV, Balagon M, Gelber R, Cho SN, Brennan PJ, Vissa V. 2009. Population-based molecular epidemiology of leprosy in Cebu, Philippines. J Clin Microbiol 47:2844–2854. 74. Cambau E, Chauffour-Nevejans A, Tejmar-Kolar L, Matsuoka M, Jarlier V. 2012. Detection of antibiotic resistance in leprosy using GenoType LepraeDR, a novel readyto-use molecular test. PLoS Negl Trop Dis 6:e1739. doi:10.1371/journal.pntd.0001739. 75. Cole ST, Eiglmeier K, Parkhill J, James KD, Thomson NR, Wheeler PR, Honoré N, Garnier T, Churcher C, Harris D, Mungall, Basham D, Brown D, Chillingworth T, Connor R, Davies RM, Devlin K, Duthoy S, Feltwell T, Fraser A, Hamlin N, Holroyd S, Hornsby T, Jagels K, Lacroix C, Maclean J, Moule S, Murphy L, Oliver K, Quail MA, Rajandream MA, Rutherford KM, Rutter S, Seeger K, Simon S, Simmonds M, Skelton J, Squares S, Stevens K, Taylor K, Whitehead S, Woodward JR, Barrell BG. 2001. Massive gene decay in the leprosy bacillus. Nature 409:1007–1011. 76. Rojas-Espinosa O, Lovik M. 2001. Mycobacterium leprae and Mycobacterium lepraemurium infections in domestic and wild animals. Rev Sci Tech 20:219–251. 77. Vera-Cabrera L, Escalante-Fuentes WG, Gomez-Flores M, Ocampo-Candiani J, Busso P, Singh P, Cole ST. 2011. Base of diffuse lepromatous leprosy associated with “Mycobacterium lepromatosis.” J Clin Microbiol 49:4366–4368. 78. Jessamine PG, Desjardins M, Gillis T, Scollard D, Jamieson F, Broukhanski G, Chedore P, McCarthy A. 2012. Leprosy-like illness in a patient with Mycobacterium lepromatosis from Ontario, Canada. J Drugs Dermatol 11:229–233. 79. Han XY, Jessurun J. 2013. Severe leprosy reactions due to Mycobacterium lepromatosis. Am J Med Sci 345:65–69. 80. Kendall BA, Winthrop KL. 2013. Update on the epidemiology of pulmonary nontuberculous mycobacterial infections. Semin Respir Crit Care Med 34:87–94. 81. Kotilainen H, Valtonen V, Tukiainen P, Poussa T, Eskola J, Järvinen A. 2013. Prognostic value of American Thoracic Society criteria for non-tuberculous mycobacterial disease: a retrospective analysis of 120 cases with four years of follow-up. Scand J Infect Dis 45:194–202. 82. Wallace RJ, Jr, Zhang Y, Brown BA, Dawson D, Murphy DT, Wilson R, Griffith D.1998. Polyclonal Mycobacterium avium complex infections in patients with nodular bronchiectasis. Am J Respir Crit Care Med 158:1235–1244. 83. Fergie JE, Milligan TW, Henderson BM, Stafford WW. 1997. Intrathoracic Mycobacterium avium complex infec-

30. Mycobacterium: General Characteristics n

84.

85.

86.

87. 88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

tion in immunocompetent children: case report and review. J Infect Dis 24:250–253. Wolinsky E. 1995. Mycobacterial lymphadenitis in children: a prospective study of 105 nontuberculous cases with long-term follow-up. Clin Infect Dis 20:954–963. Pierre-Audigier C, Ferroni A, Sermet-Gaudelus I, Le Bourgeois M, Offredo C, Vu-Thien H, Fauroux B, Mariani P, Munck A, Bingen E, Guillemot D, Quesne G, Vincent V, Berche P, Gaillard JL. 2005. Age-related prevalence and distribution of nontuberculous mycobacterial species among patients with cystic fibrosis. J Clin Microbiol 43:3467–3470. Catherinot E, Roux AL, Vibet MA, Bellis G, Ravilly S, Lemonnier L, Le Roux E, Bernède-Bauduin C, Le Bourgeois M, Herrmann JL, Guillemot D, Gaillard JL. 2013. Mycobacterium avium and Mycobacterium abscessus complex target distinct cystic fibrosis patient subpopulations. J Cyst Fibros 12:74–80. Inderlied C, Kempler C, Bermudez L. 1993. The Mycobacterium avium complex. Clin Microbiol Rev 6:266–310. Turenne CY, Wallace R, Jr, Behr MA. 2007. Mycobacterium avium in the postgenomic era. Clin Microbiol Rev 20:205–229. Feller M, Huwiler K, Stephan R, Altpeter E, Shang H, Furrer H, Pfyffer GE, Jemmi T, Baumgartner A, Egger M. 2007. Mycobacterium avium subspecies paratuberculosis and Crohn’s disease: a systematic review and meta-analysis. Lancet Infect Dis 7:607–613. Imirzalioglu C, Dahmen H, Hain T, Billion A, Kuenne C, Chakraborty T, Domann E. 2011. Highly specific and quick detection of Mycobacterium avium subsp. paratuberculosis in feces and gut tissue of cattle and humans by multiple real-time PCR assays. J Clin Microbiol 49:1843–1852. Bruijnesteijn van Coppenraet LES, de Haas PEW, Lindeboom JA, Kuijper EJ, van Soolingen D. 2008. Lymphadenitis in children in caused by Mycobacterium avium hominissuis and not related to ‘bird tuberculosis.’ Eur J Clin Microbiol Infect Dis 27:293–299. Kaevska M, Slana I, Kralik P, Reischl U, Orosova J, Holcikova A, Pavlik I. 2011. “Mycobacterium avium subsp. hominissuis” in neck lymph nodes of children and their environment examined by culture and triplex quantitative real-time PCR. J Clin Microbiol 49:167–172. Tortoli E, Rindi L, Garcia MJ, Chiaradonna P, Dei R, Garzelli C, Kroppenstedt RM, Lari N, Mattei R, Mariottini A, Mazzarelli G, Murcia MI, Nanetti A, Piccoli P, Scarparo C. 2004. Proposal to elevate the genetic variant MAC-A, included in the Mycobacterium avium complex, to species rank as Mycobacterium chimaera sp. nov. Int J Syst Evol Microbiol 54:1277–1285. Schweickert B, Goldenberg O, Richter E, Göbel UB, Petrich A, Buchholz P, Moter A. 2008. Occurrence and clinical relevance of Mycobacterium chimaera sp. nov., Germany. Emerg Infect Dis 14:1443–1446. Murcia MI, Tortoli E, Menendez MC, Palenque E, Garcia MJ. 2006. Mycobacterium colombiense sp. nov., a novel member of the Mycobacterium avium complex and description of MAC-X as a new ITS genetic variant. Int J Syst Evol Microbiol 56:2049–2054. Vuorenmaa K, Ben Salah I, Barlogis V, Chambost H, Drancourt M. 2009. Mycobacterium colombiense and pseudotuberculous lymphadenopathy. Emerg Infect Dis 15:619– 629. van Ingen J, Boeree MJ, Kösters K, Wieland A, Tortoli E, Dekhuijzen RP, van Soolingen D. 2009. Proposal to elevate Mycobacterium avium complex ITS sequevar MACQ to Mycobacterium vulneris sp. nov. Int J Syst Evol Microbiol 59:2277–2282. Ben Salah I, Cayrou C, Raoult D, Drancourt M. 2009. Mycobacterium marseillense sp. nov., Mycobacterium timo-

99.

100.

101.

102.

103.

104.

105.

106.

107.

108.

109.

110. 111.

112.

113.

563

nense sp. nov. and Mycobacterium bouchedurhonense sp. nov., novel species in the Mycobacteriun avium complex. Int J Syst Evol Microbiol 59:2803–2808. Bang D, Herlin T, Stegger M, Andersen AB, Torkko P, Tortoli E, Thomsen VO. 2008. Mycobacterium arosiense sp. nov., a slowly growing, scotochromogenic species causing osteomyelitis in an immunocompromised child. Int J Syst Evol Microbiol 58:2398–2402. Leguizamon J, Hernandez J, Murcia M-I, Soto C-Y. 2013. Identification of potential biomarkers to distinguish Mycobacterium colombiense from other mycobacterial species. Mol Cell Probes 27:46–52. Tortoli E, Tullia Simonetti M, Dionisio D, Meli M. 1994. Cultural studies on two isolates of Mycobacterium genavense from patients with acquired immunodeficiency syndrome. Diagn Microbiol Infect Dis 18:7–12. Tortoli E, Brunello F, Cagni AE, Colombrita D, Dionisio D, Grisendi L, Manfrin V, Moroni M, Passerini Tosi C, Pinsi G, Scarparo C, Tullia Simonetti M. 1998. Mycobacterium genavense in AIDS patients, report of 24 cases in Italy and review of the literature. Eur J Epidemiol 14:219–224. Thomsen VO, Dragsted UB, Bauer J, Fuursted K, Lundgren J. 1999. Disseminated infection with Mycobacterium genavense: a challenge to physicians and microbiologists. J Clin Microbiol 37:3901–3905. Hoop RK, Böttger EC, Pfyffer GE. 1996. Etiological agents of mycobacterioses in pet birds between 1986 and 1995. J Clin Microbiol 34:991–992. Saubolle MA, Kiehn TE, White MH, Rudinsky MF, Armstrong D. 1996. Mycobacterium haemophilum: microbiology and expanding clinical and geographic spectra of diseases in humans. Clin Microbiol Rev 9:435–447. Corbett EL, Hay M, Churchyard GJ, Clayton T, Williams BG, Hayes D, Mulder D, de Cock KM. 1999. Mycobacterium kansasii and M. scrofulaceum isolates from HIV-negative South African gold miners: incidence, clinical significance and radiology. Int J Tuberc Lung Dis 3:501–507. Cho JH, Yu CH, Jin MK, Kwon O, Hong KD, Choi JY, Yoon SH, Park SH, Kim CD, Kim YL. 2012. Mycobacterium kansasii pericarditis in a kidney transplant recipient: a case report and comprehensive review of the literature. Transpl Infect Dis 14:E50–E55. doi:10.1111/j.13993062.2012.00767.x. Connick E, Levi ME. 2011. Mycobacterium kansasii cutaneous abscesses occurring as immune reconstitution inflammatory syndrome. Int J Tuberc Lung Dis 15:993–994. Taillard C, Greub G, Weber R, Pfyffer GE, Bodmer T, Zimmerli S, Frei R, Bassetti S, Rohner P, Piffaretti JC, Bernasconi E, Bille J, Telenti A, Prod’hom G. 2003. Clinical implications of Mycobacterium kansasii species heterogeneity: Swiss national survey. J Clin Microbiol 41:1240– 1244. Talbot CL, Rhodes B. 2012. An atypical mycobacterial infection of the shoulder. Int J Shoulder Surg 6:64–66. Brereton AS, El Teraifi H. 2012. Mycobacterium malmoense: dissemination causes a popliteal aneurysm in a 74-year-old man. BMJ Case Rep. 2012:pii bcr/220115471. doi:10.1136/bcr12.2011.5471. Broutin V, Banuls A-L, Aubry A, Keck N, Choisy M, Bernardet J-F, Michel C, Raymond J-C, Libert C, Barnaud A, Stragier P, Portaels F, Terru D, Belon C, Dereure O, Gutierrez C, Boschiroli M-L, van de Perre P, Cambau E, Godreuil S. 2012. Genetic diversity and population structure of Mycobacterium marinum: new insights into host and environmental specificities. J Clin Microbiol 50:3627–3634. Stinear TP, Seemann T, Harrison PF, Jenkin GA, Davies JK, Johnson PD, Abdellah Z, Arrowsmith C, Chill-

564 n

114.

115.

116.

117.

118. 119.

120.

121.

122.

123.

124.

125.

126.

127.

128.

129.

BACTERIOLOGY

ingworth T, Churcher C, Clarke K, Cronin A, Davis P, Goodhead I, Holroyd N, Jagels K, Lord A, Moule S, Mungall K, Norbertczak H, Quail MA, Rabbinowitsch E, Walker D, White B, Whitehead S, Small PM, Brosch R, Ramakrishnan L, Fischbach MA, Parkhill J, Cole ST. 2008. Insights from the complete genome sequence of Mycobacterium marinum on the evolution of Mycobacterium tuberculosis. Genome Res 18:729–741. Ucko M, Colorni A. 2005. Mycobacterium marinum infections in fish and humans in Israel. J Clin Microbiol 43:892– 895. Braun E, Sprecher H, Davidson S, Kassis I. 2013. Epidemiology and clinical significance of non-tuberculous mycobacteria isolated from pulmonary specimens. Int J Tuberc Lung Dis 17:96–99. Levy I, Grisaru-Soen G, Lerner-Geva L, Kerem E, Blau H, Bentur L, Aviram M, Rivlin J, Picard E, Lavy A, Yahav Y, Rahav G. 2008. Multicenter cross-sectional study of nontuberculous mycobacterial infections among cystic fibrosis patients, Israel. Emerg Infect Dis 14:378– 384. Kent PT, Kubica GP. 1985. Public Health Mycobacteriology: A Guide for the Level III Laboratory. Centers for Disease Control, US Department of Health and Human Services, Atlanta, GA. Portaels F, Silva MT, Meyers WM. 2009. Buruli ulcer. Clin Dermatol 27:291–305. World Health Organization. 2012. Buruli Ulcer Disease. WHO Fact Sheet No. 199. World Health Organization, Geneva, Switzerland. http://www.who.int/mediacentre/ factsheets/fs199/en/. Portaels F, Chemlal K, Elsen P, Johnson PDR, Hayman JA, Hibble J, Kirkwood R, Meyers WM. 2001. Mycobacterium ulcerans in wild animals. Rev Sci Tech 20:252–264. Affolabi D, Sanoussi N, Vandelannoote, Odoun M, Faihun F, Sopoh G, Anagonou S, Portaels F, Eddyani M. 2012. Effects of decontamination, DNA extraction, and amplification procedures on the molecular diagnosis of Mycobacterium ulcerans disease (Buruli ulcer). J Clin Microbiol 50:1195–1198. Demangel C, Stinear TP, Cole ST. 2009. Buruli ulcer: reductive evolution enhances pathogenicity of Mycobacterium ulcerans. Nat Rev Microbiol 7:50–60. Doig KD, Holt KE, Fyfe JAM, Lavender CJ, Eddyani M, Portaels F, Yeboah-Manu D, Pluschke G, Seemann T, Stinear TP. 2012. On the origin of Mycobacterium ulcerans, the causative agent of Buruli ulcer. BMC Genomics 13:258–277. Grech M, Carter R, Thomson R. 2010. Clinical significance of Mycobacterium asiaticum isolates in Queensland, Australia. J Clin Microbiol 48:162–167. Lalande V, Barbut F, Vernerot A, Febvre M, Nesa D, Wadel S, Vincent V, Petit JC. 2001. Pseudo-outbreak of Mycobacterium gordonae associated with water from refrigerated fountains. J Hosp Infect 48:76–79. Pinho L, Santos J, Oliveira G, Pestana M. 2009. Mycobacterium gordonae urinary infection in a renal transplant recipient. Transpl Infect Dis 11:253–256. Verma N, Spencer D. 2012. Disseminated Mycobacterium gordonae infection in a child with cystic fibrosis. Pediatr Pulmonol 47:518–519. Eckburg PB, Buadu EO, Stark P, Sarinas PSA, Chitkara RK, Kuschner WG. 2000. Clinical and chest radiographic findings among persons with sputum culture positive for Mycobacterium gordonae. Chest 117:96–102. Tortoli E, Gitti Z, Klenk HP, Lauria S, Mannino R, Mentagani P, Mariottini A, Neonakis I. 2013. Survey of 150 strains belonging to the Mycobacterium terrae complex and description of Mycobacterium engbaekii sp. nov., Mycobacterium heraklionense sp. nov. and Mycobacterium longobardum sp. nov. Int J Syst Evol Microbiol 63:401–411.

130. Clinical and Laboratory Standards Institute (CLSI). 2008. Laboratory Detection and Identification of Mycobacteria. Document M48-A. CLSI, Wayne, PA. 131. Jensen PA, Lamber LA, Iademarco MF, Rizon R. 2005. Guidelines for preventing transmission of Mycobacterium tuberculosis in health care settings. Morbid Mortal Wkly Rep 54(RR-17):1–139. 132. Garcia LS. 2010. Clinical Microbiology Procedures Handbook, 3rd ed. ASM Press, Washington, DC. 133. Wallace R. 1987. Nontuberculous mycobacteria and water: a love affair with increasing clinical importance. Infect Dis Clin North Am 1:677–686. 134. American Thoracic Society, Centers for Disease Control and Prevention, Infectious Diseases Society of America. 2005. Controlling tuberculosis in the United States. Am J Respir Crit Care Med 172:1169–1227. 135. Mase SR, Ramsay A, Ng V, Henry M, Hopewell PC, Cunningham J, Urbanczik R, Perkins MD, Aziz MA, Pai M. 2007. Yield of serial sputum specimen examinations in the diagnosis of tuberculosis: a systematic review. Int J Tuberc Lung Dis 11:485–495. 136. Mixides G, Shende V, Teeter LD, Awe R, Musser JM, Graviss EA. 2005. Number of negative acid-fast smears needed to adequately assess infectivity of patients with pulmonary tuberculosis. Chest 128:108–115. 137. Monkongdee P, McCarthy KD, Cain KP, Tasaneepayan T, Dung NH, Lan NT, Yen NT, Teeratakulpisarn N, Udomsantisuk N, Heilig C, Varma JK. 2009. Yield of acid-fast smear and mycobacterial culture for tuberculosis diagnosis in people with HIV. Am J Respir Crit Care Med 180:903–908. 138. Noeske J, Dopico E, Torrea G, Wang H, van Deun A. 2009. Two vs. three sputum samples for microscopic detection of tuberculosis in a high HIV prevalence population. Int J Tuberc Lung Dis 13:842–847. 139. Hepple R, Ford N, McNerney R. 2012. Microscopy compared to culture for the diagnosis of tuberculosis in induced sputum samples: a systematic review. Int J Tuberc Lung Dis 16:579–588. 140. Archibald LK, McDonald LC, Addison RM, McKnight C, Byrne T, Dobbie H, Nwanyanwu O, Kezembe P, Reller LB, Jarvis WR. 2000. Comparison of Bactec MYCO/F LYTIC and WAMPOLE ISOLATOR 10 (lysiscentrifugation) systems for detection of bacteremia, mycobacteremia, and fungemia in a developing country. J Clin Microbiol 3:2994–2997. 141. Kirby JE, Delaney M, Qian Q, Gold HS. 2009. Optimal use of Myco/F Lytic and standard BACTEC blood culture bottles for detection of yeast and mycobacteria. Arch Pathol Lab Med 133:93–96. 142. Hänscheid T, Monteiro C, Melo Cristino J, Marques Lito L, Salgano MJ. 2005. Growth of Mycobacterium tuberculosis in conventional BacT/Alert FA blood culture bottles allows reliable diagnosis of mycobacteremia. J Clin Microbiol. 43:890–891. 143. Morris A, Reller LB, Salfinger M, Jackson K, Sievers A, Dwyer B. 1993. Mycobacteria in stool specimens: the nonvalue of smears for predicting culture results. J Clin Microbiol 31:1385–1387. 144. Pfyffer GE, Welscher HM, Kissling P. 1997. Pretreatment of clinical specimens with sodium dodecyl (lauryl) sulfate is not suitable for the Mycobacteria Growth Indicator Tube cultivation method. J Clin Microbiol 35:2142– 2144. 145. Padilla E, Manterola JM, Gonzalez V, Thornton CG, Quesada MD, Sanchez MD, Perez M, Ausina V. 2005. Comparison of the sodium hydroxide specimen processing method with the C18-carboxypropylbetaine specimen processing method using independent specimens with auramine smear, the MB/BacT liquid culture system, and the

30. Mycobacterium: General Characteristics n

146.

147.

148.

149.

150.

151.

152.

153.

154.

155.

156.

157.

158.

159.

COBAS AMPLICOR MTB test. J Clin Microbiol 43:6091–6097. Ferroni A, Vu-Thien H, Lanotte P, Le Bourgeois M, Sermet-Gaudelus I, Fauroux B, Marchand S, Varaigne F, Berche P, Gaillard J-L, Offredo C. 2006. Value of the chlorhexidine decontamination method for recovery of nontuberculous mycobacteria from sputum samples of patients with cystic fibrosis. J Clin Microbiol 44:2237– 2239. Bobadilla-del-Valle M, Ponce-de-Leon A, Kato-Maeda M, Hernandez-Cruz A, Calva-Mercado JJ, Chavez-Mazari B, Caballero-Rivera BA, Nolasco-Garcia JC, Sifuentes-Osornio C. 2003. Comparison of sodium bicarbonate, cetyl-pyridinium chloride, and sodium borate for preservation of sputa for culture of Mycobacterium tuberculosis. J Clin Microbiol 41:4487–4488. Cattamanchi A, Davis JL, Worodria W, Yoo S, Matovu J, Kiidha J, Nankya F, Kyeyune R, Andama A, Joloba M, Osmond D, Hopewell P, Huang L. 2008. Poor performance of universal sample processing method for diagnosis of pulmonary tuberculosis by smear microscopy and culture in Uganda. J Clin Microbiol 46:3325–3329. Palomino JC, Portaels F. 1998. Effects of decontamination methods and culture conditions on viability of Mycobacterium ulcerans in the Bactec system. J Clin Microbiol 36:402–408. Yeboah-Manu D, Bodmer T, Mensah-Quainoo E, Owusu S, Ofori-Adjei D, Pluschke G. 2004. Evaluation of decontamination methods and growth media for primary isolation of Mycobacterium ulcerans from surgical specimens. J Clin Microbiol 42:5875–5876. Burman WJ, Reves RR. 2000. Review of false-positive cultures for Mycobacterium tuberculosis and recommendations for avoiding unnecessary treatment. Clin Infect Dis 31:1390–1395. Yajko DM, Nassos PS, Sanders CA, Gonzalez PC, Reingold AL, Horsburgh CR, Hopewell P, Chin DP, Hadley WK. 1993. Comparison of four decontamination methods for recovery of Mycobacterium avium complex from stools. J Clin Microbiol 31:302–306. Lipsky BA, Gates J, Tenover FC, Plorde JJ. 1984. Factors affecting the clinical value of microscopy for acidfast bacilli. Rev Infect Dis 6:214–222. Yajko DM, Nassos PS, Sanders CA, Madej JJ, Hadley WK. 1994. High predictive value of the acid-fast smear for Mycobacterium tuberculosis despite the high prevalence of Mycobacterium avium complex in respiratory specimens. Clin Infect Dis 19:334–336. Wilmer A, Bryce E, Grant J. 2011. The role of the third acid-fast bacillus smear in tuberculosis screening for infection control purposes: a controversial topic revisited. Can J Infect Dis Med Microbiol 22(1):e1–e3. Trifiro S, Bourgault A-M, Lebel F, René P. 1990. Ghost mycobacteria on Gram stain. J Clin Microbiol 28:146– 147. Chen P, Shi M, Feng G-D, Liu J-Y, Wang B-J, Shi XD, Ma L, Liu X-D, Yang Y-N, Dai W, Liu T-T, He Y, Li J-G, Hao X-K, Zhao G. 2012. A highly efficient ZiehlNeelsen stain: identifiying de novo intracellular Mycobacterium tuberculosis and improving detection of extracellular M. tuberculosis in cerebrospinal fluid. J Clin Microbiol 50:1166–1170. Morris AJ, Reller LB. 1993. Reliability of cord formation in BACTEC media for presumptive identification of mycobacteria. J Clin Microbiol 31:2533–2534. Sanchez-Chardi A, Olivares F, Byrd TF, Julian E, Brambilla C, Luquin M. 2011. Demonstration of cord formation by rough Mycobacterium abscessus variants: implications for the clinical microbiology laboratory. J Clin Microbiol 49:2293–2295.

565

160. Attorri S, Dunbar S, Clarridge JE, III. 2000. Assessment of morphology for rapid presumptive identification of Mycobacterium tuberculosis and Mycobacterium kansasii. J Clin Microbiol 38:1426–1429. 161. Joseph S, Vaichulis E, Houk V. 1967. Lack of auraminerhodamine fluorescence of Runyon group IV mycobacteria. Am Rev Respir Dis 95:114–115. 162. Somoskövi A, Hotaling JE, Fitzgerald M, O’Donnell D, Parsons LM, Salfinger M. 2001. Lessons from a proficiency testing event for acid-fast microscopy. Chest 120:250–257. 163. Minion J, Pai M, Ramsay A, Menzies D, Greenaway C. 2011. Comparison of LED and conventional fluorescence microscopy for detection of acid fast bacilli in a lowincidence setting. PLoS ONE 6:e22495. doi:10.1371/journal.pone.0022495. 164. Habtamu M, van den Boogaard J, Ndaro A, Buretta R, Irongo CF, Lega DA, Nyombi BM, Kibiki GS. 2012. Light-emitting diode with various sputum smear preparation techniques to diagnose tuberculosis. Int J Tuberc Lung Dis 16:402–407. 165. Cuevas LE, Al-Sonboli A, Lawson L, Yassin MA, Arbide I, Al-Aghbari N, Sherchand JB, Al-Absi A, Emenyonu EN, Merid Y, Okobi MI, Onuoha JO, Aschalew M, Aseffa A, Harper G, Anderson de Cuevas RM, Theobald SJ, Nathanson C-M, Joly J, Faragher B, Squire SB, Ramsay A. 2011. LED fluorescence microscopy for the diagnosis of pulmonary tuberculosis: a multi-country cross-sectional evaluation. PLoS Med 8:e1001057. doi:10.1371/journal.pmed.1001057. 166. Xia H, Song Y-Y, Zhao B, Kam K-M, O’Brien RJ, Zhang Z-Y, Sohn H, Wang W, Zhao Y-L. 2013. Multicentre evaluation of Ziehl-Neelsen and light-emitting diode fluorescence microscopy in China. Int J Tuberc Lung Dis 17:107–112. 167. Minion J, Shenai S, Vadwai V, Tipnis T, Greenaway C, Menzies D, Ramsay A, Rodrigues C, Pai M. 2011. Fading of auramine-stained mycobacterial smears and implications for external quality assurance. J Clin Microbiol 49:2024–2026. 168. Murray SJ, Barrett A, Magee JG, Freeman R. 2003. Optimisation of acid-fast smears for the direct detection of mycobacteria in clinical samples. J Clin Pathol 56:613– 615. 169. European Society of Mycobacteriology. 1991. Decontamination, microscopy and isolation, p 57–64. In Yates MD, Groothuis DG (ed), Diagnostic Public Health Mycobacteriology. Bureau of Hygiene and Tropical Disease, London, United Kingdom. 170. Warren JR, Bhattacharya M, de Almeida KN, Trakas K, Peterson LR. 2000. A minimum 5.0 ml of sputum improves the sensitivity of acid-fast smear for Mycobacterium tuberculosis. Am J Respir Crit Care Med 161:1559– 1562. 171. Saceanu C, Pfeiffer N, McLean T. 1993. Evaluation of sputum smears concentrated by cytocentrifugution for detection of acid-fast bacilli. J Clin Microbiol 31:2371– 2374. 172. Woods GL, Pentony E, Boxley MJ, Gatson AM. 1995. Concentration of sputum by cytocentrifugation for preparation of smears for detection of acid-fast bacilli does not increase sensitivity of the fluorochrome stain. J Clin Microbiol 33:1915–1916. 173. Yeboah-Manu D, Danso E, Ampah K, Asante-Poku A, Nakobu Z, Pluschke G. 2011. Isolation of Mycobacterium ulcerans from swab and fine-needle-aspiration specimens. J Clin Microbiol 49:1997–1999. 174. Welch D, Guruswamy A, Sides S, Shaw C, Gilchrist M. 1993. Timely culture of mycobacteria which utilizes a microcolony method. J Clin Microbiol 31:2178–2184.

566 n

BACTERIOLOGY

175. Clinical and Laboratory Standards Institute (CLSI). 2009. Susceptibility Testing of Mycobacteria, Nocardiae, and Other Aerobic Actinomycetes. Document M24-A2. CLSI, Wayne, PA. 176. Leung E, Minion J, Benedetti A, Pai M, Menzies D. 2011. Microcolony culture techniques of tuberculosis diagnosis: a systematic review. Int J Tuberc Lung Dis 16:16– 23. 177. Realini L, de Ridder K, Hirschel B, Portaels F. 1999. Blood and charcoal added to acidified agar media promote growth of Mycobacterium genavense. Diagn Microbiol Infect Dis 34:45–50. 178. Saito H, Toda K, Matsumoto I, Matsuo K, Nakanaga K, Ishii N. 2004. Bacteriological features of Mycobacterium haemophilum isolated from skin lesions in an immunodeficient patient. Kansenshogaku Zasshi 78:389–397. 179. Piersimoni C, Scarparo C, Callegaro A, Passerini Tosi C, Nista D, Bornigia S, Scagnelli M, Rigon A, Ruggiero G, Goglio A. 2001. Comparison of MB/BacT Alert 3D System with radiometric Bactec System and LöwensteinJensen medium for recovery and identification of mycobacteria from clinical specimen: a multicenter study. J Clin Microbiol 39:651–657. 180. Pfyffer GE, Welscher HM, Kissling P, Cieslak C, Casal MJ, Gutierrez J, Rüsch-Gerdes S. 1997. Comparison of the Mycobacteria Growth Indicator Tube (MGIT) with radiometric and solid culture for recovery of acid-fast bacilli. J Clin Microbiol 35:364–368. 181. Whyte T, Hanahoe B, Collins T, Corbett-Feeney G, Cormican M. 2000. Evaluation of the Bactec MGIT 960 and MB/BacT Systems for routine detection of Mycobacterium tuberculosis. J Clin Microbiol 38:3131–3132. 182. Gravet A, Souillard N, Habermacher J, Moser A, Lohmann C, Schmitt F, Delarbre JM. 2011. Culture and susceptibility testing of mycobacteria with VersaTREK. Pathol Biol (Paris) 59:32–38. 183. Cruciani M, Scarpaio C, Malena M, Bosco O, Serpelloni G, Mengoli C. 2004. Meta-analysis of Bactec MGIT 960 and Bactec 460 TB, with or without solid media, for detection of mycobacteria. J Clin Microbiol 42:2321–2325. 184. Parrish N, Dionne K, Sweeney A, Hedgepeth A, Carroll K. 2009. Differences in time to detection and recovery of Mycobacterium spp. between the MGIT 960 and the BacT/ALERT MB automated culture systems. Diagn Microbiol Infect Dis 63:342–345. 185. Crump JA, Morrissey AB, Ramadhani HO, Njau BN, Maro VP, Reller LB. 2011. Controlled comparison of BacT/Alert MB System, manual Myco/F Lytic procedure, and Isolator 10 System for diagnosis of Mycobacterium tuberculosis bacteremia. J Clin Microbiol 49:3054–3057. 186. Pena JA, Ferraro MJ, Hoffmann CG, Branda JA. 2012. Growth detection failures by the non-radiometric Bactec MGIT 960 mycobacterial culture system. J Clin Microbiol 50:2092–2095. 187. Tyrrell FC, Budnick GE, Elliott T, Gillim-Ross L, Hildred MV, Mahlmeister P, Parrish N, Pentella M, Vanneste J, Wang YF, Starks AM. 2012. Probability of negative Mycobacterium tuberculosis complex cultures based on time to detection of positive cultures: a multicenter evaluation of commercial-broth-based culture systems. J Clin Microbiol 50:3275–3282. 188. Pfyffer GE, Wittwer F. 2012. Incubation time of mycobacterial cultures: how long is long enough to issue a final negative report to the clinician? J Clin Microbiol 50:4188– 4189. 189. Shu Z, Weigel KM, Soelberg SD, Lakey A, Cangelosi GA, Lee K-H, Chung J-H, Gao D. 2012. Cryopreservation of Mycobacterium tuberculosis complex cells. J Clin Microbiol 50:3573–3580. 190. Nienhaus A, Schablon A, Costa JT, Diel R. 2011. Systematic review of cost and cost-effectiveness of different

191.

192.

193.

194.

195.

196.

197.

198.

199.

200.

201.

202.

203.

204.

TB-screening strategies. BMC Health Serv Res 11:247– 257. Gandra S, Scott WS, Somaraju V, Wang H, Wilton S, Feigenbaum M. 2010. Questionable effectiveness of the QuantiFERON-TB Gold Test (Cellestis) as a screening tool in healthcare workers. Infect Control Hosp Epidemiol 31:1279–1285. Detjen AK, Loebenberg L, Grewal HM, Stanley K, Gutschmidt A, Kruger C, Du Plessis N, Kidd M, Beyers N, Walzl G, Hesseling AC. 2009. Short-term reproducibility of a commercial interferon gamma release assay. Clin Vaccine Immunol 16:1170–1175. Centers for Disease Control and Prevention. 2010. Updated guidelines for using interferon gamma release assays to detect Mycobacterium tuberculosis infection—United States 2010. Morbid Mortal Wkly Rep 59:1–26. Zellweger J-P, Zellweger A, Ansermet S, de Senarclens B, Wrighton-Smith P. 2005. Contact tracing using a new T-cell based test: better correlation with tuberculosis exposure than the tuberculin test. Int J Tuberc Lung Dis 9:1242–1247. Pai M, Zwerling A, Menzies D. 2008. Systematic review: T-cell-based assays for the diagnosis of latent tuberculosis infection: an update. Ann Intern Med 149:177–184. Higuchi K, Sekiya Y, Igari H, Watanabe A, Harada N. 2012. Comparison of specificities between two interferongamma release assays in Japan. Int J Tuberc Lung Dis 16:1190–1192. Diel R, Goletti D, Ferrara G, Bothamley G, Cirillo D, Kampmann B, Lange C, Losi M, Markova R, Migliori GB, Nienhaus A, Ruhwald M, Wagner D, Zellweger JP, Huitric E, Sandgren A, Manissero D. 2011. Interferonγ release assays for the diagnosis of latent Mycobacterium tuberculosis infection: a systematic review and metaanalysis. Eur Respir J 37:88–99. Sester M, Sotgiu G, Lange C, Giehl C, Girardi E, Migliori GB, Bossink A, Dheda K, Diel R, Dominguez J, Lipman M, Nemeth J, Ravn P, Winkler S, Huitric E, Sandgren A, Manissero D. 2011. Interferon-γ release assays for the diagnosis of active tuberculosis: a systematic review and meta-analysis. Eur Respir J 37:100–111. Chen J, Zhang R, Wang J, Liu L, Zheng Y, Shen Y, Qi T, Lu H. 2011. Interferon-gamma release assays for the diagnosis of active tuberculosis in HIV-infected patients: a systematic review and meta-analysis. PLoS One 6(11):e26827. doi:10.1371/journal.pone.0026827. Denkinger CM, Pai M, Patel M, Menzies D. 2013. Gamma interferon release assay for monitoring of treatment response for active tuberculosis: an explosion in the spaghetti factory. J Clin Microbiol 51:607–610. Diel R, Loddenkemper R, Meywald-Walter K, Gottschalk R, Nienhaus A. 2009. Comparative performance of tuberculin test, QuantiFERON-TB-Gold In Tube assay, and T-Spot.TB test in contact investigations for tuberculosis. Chest 135:1010–1018. Stephan C, Wolf T, Goetsch U, Bellinger O, Nisius G, Oremek G, Rakus Z, Gottschalk R, Stark S, Brodt HR, Staszewski S. 2009. Comparing QuantiFERON-tuberculosis Gold, T-SPOT tuberculosis and tuberculin skin test in HIV-infected individuals from a low prevalence tuberculosis country. AIDS 22:2471–2479. Scrivo R, Sauzullo I, Mengoni F, Priori R, Coppola M, Iaiani G, Di Franco M, Vullo V, Mastroianni CM, Valesini G. 2013. Mycobacterial interferon-release variations during longterm treatment with tumor necrosis factor blockers: lack of correlation with clinical outcome. J Rheumatol 40:157–165. Tavast E, Salo E, Seppälä I, Tuuminen T. 2009. IGRA tests perform similarly to TST but cause no adverse reactions: pediatric experience in Finland. BMC Res Notes 2:9–18.

30. Mycobacterium: General Characteristics n 205. Kampmann B, Whittaker E, Williams A, Walters S, Gordon A, Martinez-Alier N, Williams B, Crook AM, Hutton AM, Anderson ST. 2009. Interferon-gamma release assays do not identify more children with active TB than TST. Eur Respir J 33:1374–1382. 206. Mandalakas AM, Detjen AK, Hesseling AC, Benedetti A, Menzies D. 2011. Interferon-gamma release assays and childhood tuberculosis: systematic review and metaanalysis. Int J Tuberc Lung Dis 15:1018–1032. 207. Zellweger J-P. 2008. Latent tuberculosis: which test in which situation? Swiss Med Wkly 138:31–37. 208. Menzies D, Pai M, Comstock G. 2007. Meta-analysis: new tests for the diagnosis of latent tuberculosis infection. Areas of uncertainty and recommendations for research. Ann Intern Med 146:340–354. 209. Chee CBE, Gan SH, Khinmar KW, Barkham TM, Koh CK, Liang S, Wang YT. 2008. Comparison of sensitivities of two commercial gamma interferon release assays for pulmonary tuberculosis. J Clin Microbiol 46:1935–1940. 210. Pai M, Joshi R, Dogra S, Zwerling AA, Gajalakshmi D, Goswami K, Reddy MVR, Kalantri S, Hill PC, Menzies D, Hopewell PC. 2009. T-cell assay conversions and reversions among household contacts of tuberculosis patients in rural India. Int J Tuberc Lung Dis 13:84–92. 211. Pai M, O’Brien R. 2007. Serial testing for tuberculosis: can we make sense of T cell assay conversions and reversions? PLoS Med 4:980–983. 212. Shah M, Kasambira TS, Adrian PV, Madhi SA, Martinson NA, Dorman SE. 2011. Longitudinal analysis of QuantiFERON-TB Gold In-Tube in children with adult household tuberculosis contact in South Africa: a prospective cohort study. PLoS One 6:e26787. doi:10.1371/ journal.pone.0026787. 213. van Zyl-Smit RN, Pai M, Peprah K, Meldau R, Kieck J, Jurizu J, Badri M, Zumla A, Sechi LA, Bateman ED, Dheda K. 2009. Within-subject variability and boosting of T-cell interferon-γ responses after tuberculin skin testing. Am J Respir Crit Care Med 180:49–58. 214. Talati NJ, Seybold U, Humphrey B, Aina A, Tapia J, Weinfurter P, Albalak R, Blumberg HM. 2009. Poor concordance between interferon-gamma release assays and tuberculin tests in diagnosis of latent tuberculosis infection among HIV-infected individuals. BMC Infect Dis 9:15–24. 215. Bergamini BM, Losi M, Valenti F, D’Amico R, Meccugni B, Meacci M, de Giovanni D, Rumianesi F, Fabbri LM, Balli F, Richeldi L. 2009. Performance of commercial blood tests for the diagnosis of latent tuberculosis infection in children and adolescents. Pediatrics 123:e419–e424. 216. Kobashi Y, Sugiu T, Shimizu H, Ohue Y, Mouri K, Obase Y, Miyashita N, Oka M. 2009. Clinical evaluation of the T-SPOT.TB Test for patients with indeterminate results on the QuantiFERON TB-2G Test. Intern Med 48:137–142. 217. Banach DB, Harris TG. 2011. Indeterminate QuantiFERON-TB Gold results in a public health clinic setting. Int J Tuberc Lung Dis 15:1623–1629. 218. Herrera V, Yeh E, Murphy K, Parsonnet J, Banaei N. 2010. Immediate incubation reduces indeterminate results for QuantiFERON TB Gold In-Tube assay. J Clin Microbiol 48:2672–2676. 219. Doberne D, Gaur RL, Banaei N. 2011. Preanalytical delay reduces sensitivity of QuantiFERON-TB Gold InTube Assay for detection of latent tuberculosis infection. J Clin Microbiol 49:3061–3064. 220. Centers for Disease Control and Prevention. 1997. Multiple misdiagnoses of tuberculosis resulting from laboratory error—Wisconsin 1996. Morbid Mortal Wkly Rep 46:797–801.

567

221. Killeen AA. 2009. Laboratory sanctions for proficiency testing sample referral and result communication: a review of actions from 1993–2006. Arch Pathol Lab Med 133:979–982. 222. Hawkins JE, Good RC, Kubica GP, Gangadharam PRJ, Gruft HM, Stottmeier KD, Sommers HM, Wayne LG. 1983. Levels of laboratory services for mycobacterial diseases: official statement of the American Thoracic Society. Am Rev Respir Dis 128:213. 223. Woods GL, Ridderhof JC. 1996. Quality assurance in the mycobacteriology laboratory. Clin Lab Med 16:657– 675. 224. Clinical and Laboratory Standards Institute (CLSI). 2006. Molecular Diagnostic Methods for Infectious Diseases. Document MM03-A2. CLSI, Wayne, PA. 225. Centers for Disease Control and Prevention. 2013. Availability of an assay for detecting Mycobacterium tuberculosis, including rifampin-resistant strains, and considerations for its use—United States, 2013. Morbid Mortal Wkly Rep 62:821–824. 226. Masaki T, Ohkusu K, Hata H, Fujiwara N, Iihara H, Yamada-Noda M, Nhung PH, Hayashi M, Asano Y, Kawamura Y, Ezaki T. 2006. Mycobacterium kumamotonense sp. nov., recovered from clinical specimen and the first isolation report of Mycobacterium arupense in Japan: novel slowly growing, non-chromogenic clinical isolates related to Mycobacterum terrae complex. Microbiol Immunol 50:889–897. 227. Tsai TF, Lai CC, Chang CH, Hsiao CH, Hsueh PR. 2006. Tenosynovitis caused by Mycobacterium arupense in a patient with diabetes mellitus. Clin Infect Dis 47:861– 863. 228. Huber J, Richter E, Binder L, Maass M, Eberl R, Zenz W. 2008. Mycobacterium bohemicum and cervical lymphadenitis. Emerg Infect Dis 14:1158–1159. 229. Poulin S, Corbeil C, Nguyen M, St-Denis A, Coté L, Le Deist F, Carignan A. 2013. Fatal Mycobacterium colombiense/cytomegalovirus coinfection associated with acquired immunodeficiency due to autoantibodies against interferon gamma: a case report. BMC Infect Dis 13:24–29. 230. van Ingen J, Boeree MJ, Stals FS, Pitz CCM, RooijmansRietjens JJCM, van der Zanden AGM, Dekhuijzen PNR, van Soolingen D. 2007. Clinical Mycobacterium conspicuum isolation from two immunocompetent patients in the Netherlands. J Clin Microbiol 45:4075–4076. 231. Tortoli E, Piersimoni C, Kroppenstedt RM, MontoyaBurgos JI, Reischl U, Giacometti A, Emler S. 2001. Mycobacterium doricum sp. nov. Int J Syst Evol Microbiol 51:2007–2012. 232. Pettit AC, Jahangir AA, Wright PW. 2011. Mycobacterium doricum osteomyelitis and soft tissue infection. Emerg Infec Dis 17:2075–2076. 233. Tortoli E, Böttger E, Fabio A, Falsen E, Gitti Z, Grottola A, Klenk HP, Mannino R, Mariottini A, Messino M, Pecorari M, Rumpianesi F. 2011. Mycobacterium europaeum sp. nov., a scotochromogenic species related to the Mycobacterium simiae complex. Int J Syst Evol Microbiol 61:1606–1611. 234. Tortoli E, Rindi L, Goh KS, Katila ML, Mariottini A, Mattei R, Mazzarelli G, Suomalainen S, Torkko P, Rastogi N. 2005. Mycobacterium florentinum sp. nov., isolated from humans. Int J Syst Evol Microbiol 55:1101–1106. 235. Ramos JP, Campos CE, Caldas PC, Ferreira NV, da Silva MV, Redner P, Campelo CL, Vale SF, Barroso EC, Medeiros RF, Montes FC, Galvao TC, Tortoli E. 2013. Mycobacterium fragae sp. nov., a non-chromogenic species isolated from human respiratory specimens. Int J Syst Evol Microbiol 63:2583–2587. 236. Okazaki M, Ohkusu K, Hata H, Ohnisi H, Sugahara K, Kawamura C, Fujiwara N, Matsumoto S, Nishiuchi

568 n

237.

238.

239.

240.

241.

242.

243.

244.

245.

246.

247.

248.

249.

BACTERIOLOGY

Y, Toyoda K, Saito H, Yonetani S, Fukugawa Y, Yamamoto M, Wada H, Sejimo A, Ebina A, Goto H, Ezaki T, Watanabe T. 2009. Mycobacterium kyorinense sp. nov., a novel slow-gowing species, related to Mycobacterium celatum, isolated from human clinical specimens. Int J Syst Evol Microbiol 59:1336–1341. Dias Campos CE, de Souza Caldas PC, Ohnishi H, Watanabe T, Ohtsuka K, Matsushima S, Ferreira NV, Villas Boas da Silva M, Redner P, Distasio de Carvalho L, Ferreira de Melo Medeiros R, Abbud Filho JA, Onofre Fandinho Montes C, Calcagno Galvao T, Pais Ramos J. 2012. First isolation of Mycobacterium kyorinense from clinical specimens in Brazil. J Clin Microbiol 50:2477–2478. Hong SK, Sung JY, Lee HJ, Oh MD, Park SS, Kim EC. 2013. First case of Mycobacterium longobardum infection. Ann Lab Med 33:356–359. van Ingen J, Lindenboom JA, Hartwig NG, de Zwaan RE, Tortoli E, Dekhuijzen RP, Boeree MJ, van Soolingen D. 2009. Mycobacterium mantenii sp. nov., a pathogenic, slowly growing, scotochromogenic mycobacterium. Int J Syst Evol Microbiol 59:2782–2787. Mohamed AM, Iwen PC, Tarantolo S, Hinrichs SH. 2004. Mycobacterium nebraskense sp. nov., a novel slowly growing scotochromogenic species. Int J Syst Evol Microbiol 54:2057–2060. Turenne CY, Cook VJ, Burdz TV, Pauls RJ, Thibert L, Wolfe JN, Kabani A. 2004. Mycobacterium parascrofulaceum sp. nov., novel slowly growing, scotochromogenic clinical isolates related to Mycobacterium simiae. Int J Syst Evol Microbiol 54:1543–1551. Lee HK, Lee S-A, Lee I-K, Yu H-K, Park Y-G, Hyun J-W, Kim K, Kook Y-H, Kim B-J. 2010. Mycobacterium paraseoulense sp. nov., a slowly growing, scotochromogenic species related genetically to Mycobacterium seoulense. Int J Syst Evol Microbiol 60:439–443. Fanti F, Tortoli E, Hall L, Roberts GD, Kroppenstedt RM, Dodi I, Conti S, Polonelli L, Chezzi C. 2004. Mycobacterium parmense sp. nov. Int J Syst Evol Microbiol 54:1123–1127. van Ingen J, Al-Hajoj SA, Boeree M, Al-Rabiah F, Enaimi M, de Zwaan R, Tortoli E, Dekhuijzen, van Soolingen D. 2009. Mycobacterium riyadhense sp. nov., a non-tuberculous species identified as Mycobacterium tuberculosis complex by a commercial line-probe assay. Int J Syst Evol Microbiol 59:1049–1053. Godreuil S, Marchandin H, Michon A-L, Ponsada M, Chyderiotis G, Brisou P, Bhat A, Panteix G. 2012. Mycobacterium riyadhense pulmonary infection, France and Britain. Emerg Infect Dis 18:176–178. Turenne CY, Thibert L, Williams K, Burdz TV, Cook VJ, Wolfe JN, Cockcroft DW, Kabani A. 2004. Mycobacterium saskatchewanense sp. nov., a novel slowly growing scotochromogenic species from human clinical isolates related to Mycobacterium interjectum and Accuprobepositive for Mycobacterium avium complex. Int J Syst Evol Microbiol 54:659–667. van Ingen J, Tortoli E, Selvarangan R, Coyle MB, Crump JA, Morrissey AB, Dekhuijzen PNR, Boeree MJ, van Soolingen D. 2011. Mycobacterium sherrisii sp. nov., a slow-growing non-chromogenic species. Int J Syst Evol Microbiol 61:1293–1298. Selvarangan R, Wu W-K, Nguyen TT, Carlson LDC, Wallis CK, Stiglich SK, Chen Y-C, Jost, KC, Jr, Prentice JL, Wallace, RJ, Jr, Barrett SL, Cookson BT, Coyle MB. 2004. Characterization of a novel group of mycobacteria and proposal of Mycobacterium sherrisii sp. nov. J Clin Microbiol 42:52–59. Tajan J, Spasa M, Sala M, Navarro M, Font B, GonzalezMartin J, Segura F. 2013. Disseminated infection by My-

250.

251.

252.

253.

254.

255.

256.

257.

258.

259.

260.

261.

262.

cobacterium sherrisii and Histoplasma capsulatum in an African HIV-infected patient. Am J Trop Med Hyg 88:914– 917. Mun HS, Park JH, Kim H, Yu HK, Park YG, Cha CY, Kook YH, Kim BJ. 2008. Mycobacterium senuense sp. nov., a slowly growing, nonchromogenic species closely related to the Mycobacterium terrae complex. Int J Syst Evol Microbiol 58:641–646. Mun HS, Kim HJ, Oh EJ, Kim H, Bai GH, Yu HK, Park YG, Cha CY, Kook YH, Kim BJ. 2007. Mycobacterium seoulense sp. nov., a slowly growing scotochromogenic species. Int J Syst Evol Microbiol 57:594–599. Nakanaga K, Hoshino Y, Wakabayashi M, Fujimoto N, Tortoli E, Makino M, Tanaka T, Ishii N. 2012. Mycobacterium shigaense sp. nov., a novel slowly growing scotochromogenic mycobacterium that produced nodules in an erythroderma patient with severe cellular immunodeficiency and a history of Hodgkin’s disease. J Dermatol 39:389–396. Palmore TN, Shea YR, Conville PS, Witebsky FG, Anderson VL, Rupp Hodge IP, Holland SM. 2009. “Mycobacterium tilburgii,” a newly described, uncultivated opportunistic pathogen. J Clin Microbiol 47:1585–1587. Hartwig NG, Warris A, van de Vosse E, van der Zanden AGM, Schülin-Casinato T, van Ingen J, van Hest R. 2011. “Mycobacterium tilburgii” infection in two immunocompromized children: importance of molecular tools in culture-negative mycobacterial disease diagnosis. J Clin Microbiol 49:4409–4411. Kim BJ, Math RK, Jeon CO, Yu HK, Park YG, Kook YH, Kim BJ. 2013. Mycobacterium yongonense sp. nov., a slow-growing non-chromogenic species closely related to Mycobacterium intracellulare. Int J Syst Evol Microbiol 63:192–199. Tortoli E, Mariottini A, Pierotti P, Simonetti TM, Rossolini GM. 2013. Mycobacterium yongonense in pulmonary disease, Italy. Emerg Infect Dis 19:1902–1904. Federal Government. 2014. CFR, Title 42—Public Health. Chapter I—Public Health Service, Department of Health and Human Services. Subchapter F—Quarantine, Inspection, Licensing. Part 72—Interstate Transportation of Etiologic Agents. U.S. Federal Government, Washington, DC. http://www.gpo.gov/fdsys/granule/CFR-2007-title42vol1/CFR-2007-title42-vol1-part72. Federal Government. 2014. CFR, Title 49—Transportation. Chapter I—Pipeline and Hazardous Materials Safety Administration, Department of Transportation. Subchapter C—Hazardous Materials Regulations. Parts 171 to 180. U.S. Federal Government, Washington, DC. http://www .ecfr.gov/cgi-bin/text-idx?SID=c5250b44293a9c4f37c d4ec0e9f7afe1&tpl=/ecfrbrowse/Title49/49CIsubchapC .tpl. Federal Government. 2014. CFR, Title 39—Postal Service. Chapter I—United States Postal Service. Subchapter B— International Mail. Part 20—International Postal Service. U.S. Federal Government, Washington, DC. http://www .ecfr.gov/cgi-bin/text-idx?c=ecfr&tpl=/ecfrbrowse/Title 39/39cfr20_main_02.tpl. Federal Government. 2014. CFR, Title 39, Part 111. General Information on Postal Service. U.S. Federal Government, Washington, DC. http://www.ecfr.gov/cgi-bin/text -idx?rgn=div5&node=39:1.0.1.3.12. U.S. Postal Service. 2014. 346. Toxic substances and infectious substances (hazard class 6), in Publication 52 - Hazardous, Restricted, and Perishable Mail. U.S. Postal Service, Washington, DC. http://pe.usps.com/text/pub52/ pub52c3_023.htm. U.S. Postal Service. 2014. International Mail Manual, Section 135, Dangerous Mailable Goods. U.S. Postal Serv-

30. Mycobacterium: General Characteristics n ice, Washington, DC. http://pe.usps.com/text/imm/im mc1_014.htm. 263. Occupational Safety and Health Administration (OSHA). 2014. 29 CFR, Part 1910. Occupational Safety and Health Standards. Subpart Z, Toxic and Hazardous Substances. Standard 1030, Bloodborne Pathogens. Department of Labor, U.S. Federal Government, Washington, DC. https://www.osha.gov/pls/oshaweb/owadisp.show_ document?p_id=10051&p_table=STANDARDS.

569

264. International Civil Aviation Organization (ICAO). 2013–2014. Technical Instructions for the Safe Transport of Dangerous Goods by Air. Document 9284. ICAO, Mexico City, Mexico. http://www.icao.int/safety/Dangerous Goods/Pages/technical-instructions.aspx. 265. International Air Transport Association (IATA). 2014. Dangerous Goods Regulations (DGR), 55th ed. IATA, Montreal, Canada. http://www.iata.org/publications/dgr/ Pages/index.aspx.

Mycobacterium: Laboratory Characteristics of Slowly Growing Mycobacteria* PATRICIA J. SIMNER, STEFFEN STENGER, ELVIRA RICHTER, BARBARA A. BROWN-ELLIOTT, RICHARD J. WALLACE, JR., AND NANCY L. WENGENACK

31 This chapter represents a transition from previous chapters that detailed the general characteristics and phenotypic methods of identification of slowly growing mycobacteria. Although phenotypic characterization remains important, molecular techniques are increasingly employed as the gold standard for definitive identification to the species level. As Tortoli noted in 2003, mycobacterial taxonomy can be divided into two major periods defined by methods used for identification to the species level (1). The first period, characterized by utilization of phenotypic studies, lasted from the late 1880s to the end of the 1980s. The second major era, characterized by a shift to genotypic studies, began during the last decade of the 20th century and has continued to the present time. The Mycobacterium tuberculosis complex (MTBC) remains the most important group within the genus Mycobacterium from a global and clinical perspective. The MTBC currently includes not only the most significant human mycobacterial pathogens, M. tuberculosis, M. bovis, and M. bovis bacillus Calmette-Guérin (BCG), but also the less frequently encountered pathogens M. caprae, M. microti, M. africanum, M. canettii (the smooth variant of M. tuberculosis), and M. pinnipedii. Further discussion regarding the MTBC can be found in chapter 30 in this Manual. Of the more than 150 currently validated species of nontuberculous mycobacteria (NTM), approximately 80 are slowly growing species (Table 1). The most clinically significant and/or most frequently encountered slowly growing NTM species include Mycobacterium avium, M. intracellulare, M. kansasii, M. marinum, M. xenopi, M. malmoense, and M. ulcerans. M. gordonae, although rarely a pathogen, occurs frequently in human samples, usually as a consequence of contamination from tap water (Table 2). Like the rapidly growing mycobacterial (RGM) species (see chapter 32 in this Manual), the majority of species of the slowly growing NTM have been described since the early 1990s with the advent of molecular technology. From 2007 to mid-2013, approximately 25 new species were described, including M. europaeum, M. koreense, M. longobardum, M. minnesotense,

M. parakoreense, M. sherrisii, M. shinjukuense, and M. yongonense (2–23).

EPIDEMIOLOGY AND TRANSMISSION Unlike M. tuberculosis, for which humans are the definitive host, most species of NTM are widely distributed in the environment, and the occurrence of NTM disease is attributed to a combination of host factors, such as age, body weight, the presence of chronic lung diseases (such as cystic fibrosis, bronchiectasis, or chronic obstructive pulmonary disease), alterations of chest structure, and other conditions, along with exposure. Organisms can be found in samples of soil and water, including both natural and treated water sources. For example, M. kansasii, M. xenopi, and M. simiae are regularly recovered from municipal water and only rarely from other environmental sources. Furthermore, unlike MTBC, there has been no evidence of animal-to-human (except M. marinum from fish/fish tanks) or human-to-human transmission with slowly growing NTM. Human disease due to NTM is assumed to be acquired from environmental sources either directly by inhaling organisms in aerosols or traumatic implantation or indirectly by ingesting contaminated food or water. The source of infection may not always be detected for NTM (24). Incidence rates of NTM disease are only estimates since, unlike tuberculosis (TB), NTM disease is noncommunicable from human to human, and therefore numbers of infections are not required to be reported to public health agencies. One publication from 2007 reported that the isolation prevalence of all NTM species (excluding M. gordonae) in pulmonary disease in Ontario, Canada, increased from 9.1/100,000 in 1997 to 14.1/100,000 by 2003, with a mean annual increase of 8.4%. Similar increases were noted for individual species. These findings indicate a significant rise in pulmonary disease caused by NTM in Canada (25). Increasing numbers of NTM isolates were also reported for several European countries (26). Although the data are scarce, NTM disease prevalence is believed to be increasing in the United States as well (24). Nationwide studies by the Centers for Disease Control and Prevention (CDC) have revealed that NTMs

*This chapter contains information presented in chapter 29 by Elvira Richter, Barbara A. Brown-Elliott, and Richard J. Wallace, Jr., in the 10th edition of this Manual.

doi:10.1128/9781555817381.ch31

570

31. Characteristics of Slowly Growing Mycobacteria n 571 TABLE 1

Characteristics of currently recognized slowly growing species of NTMh

Species M. algericum M. arosiense M. arupense M. asiaticum M. avium subsp. avium “M. avium subsp. hominissuis” M. avium subsp. paratuberculosis M. bohemicum M. botniense M. bouchedurhonense M. branderi M. celatum M. chimaera M. colombiense M. conspicuum M. cookii M. doricum M. engbackii M. europaeum M. farcinogenes M. florentinum M. fragae M. gastri M. genavense M. gordonae M. haemophilum M. heckeshornense M. heidelbergense M. heraklionense M. hiberniae M. interjectum M. intermedium M. intracellulare M. kansasii M. kubicae M. koreense M. kumamotonense M. kyorinense M. lacus M. lentiflavum M. leprae M. lepraemurium M. longobardum M. malmoense M. mantenii M. marinum M. marseillense M. minnesotense M. monteofiorense M. nebraskense M. nonchromogenicum M. noviomagense M. palustre M. paraffinicum M. paragordonae M. parakoreense M. parascrofulaceum M. paraseoulense M. paraterrae M. parmense M. pseudoshottsii M. pulveris

Pigmentation

Established pathogenicity

Unique hsp65 gene

Unique 16S rRNA gene

N Y Nb Y Nb Nb Nb Y Y N N N N Y Y Y Y Yg Y Y N N N N Y N Y N N Y Y Y Y/N Y Y N N N N Y —h — N N Y Y N Y N Y N N Y Y Y N Y Y Y Y Y N

Ya Y Y Y Yc Ye Y Y N Y Y Y Y Y Y N Y N Y Q Y U N Y N Y Y Y Y N Y Q Y Y N Y Q Q Q Y Y Y Y Y Y Y Y N Yi Q Q N Q U N Y Y U Y Y Yj N

Y Y Y Y Yd Yd Yd U U Y U Y N N U U U Y Y Y U Y Y U U Y Y Y Y Y Y Y Y Y U Y Y Y Y Y Y Y Y Y Y Y Y Y Y Y N Y U Y Y Y Y Y Y Y N U

Y Y Y Y N N N Y Y Y Y Y Y Y Y Y Y Y Y Y Y Y N Y V Y Y Y Y Y Y Y Y N Y Y Y Y Y Y Y Y Y Y Y N Y Y Y Y Q Y Y Y Y Y Y N Y Y N Y

Date of description

2011 2008 2006 1971 1901 2002 1900 1998 2000 2009 1995 1993 2004 2006 1996 1990 2001 2013 2011 1973 2005 2013 1966 1993 1962 1978 2001 1998 2013 1993 1993 1993 1965 1955 2000 2012 2006 2009 2002 1996 1880 1912 2013 1977 2009 1926 2009 2013 2003 2004 1965 2009 2002 2009 2014 2012 2004 2010 2010 2004 2005 1983 (Continued on next page)

572 n

TABLE 1

BACTERIOLOGY

Characteristics of currently recognized slowly growing species of NTMh (Continued)

Species M. M. M. M. M. M. M. M. M. M. M. M. M. M. M. M. M. M. M. M. M. M.

riyadhense saskatchewanense scrofulaceum senuense seoulense sherrisii shigaense shimoidei shinjukuense shottsii simiae stomatepiae szulgai terrae timonense triplex triviale tusciae ulcerans vulneris xenopi yongonense

Pigmentation

Established pathogenicity

Unique hsp65 gene

Unique 16S rRNA gene

Date of description

N Y Y N Y N Y N N N Y N Y N N N N Y N Y N N

Y Y Y Q Q Y Y Y Y Yj Y Yj Y Q Y Y N Y Y Q Y Y

Y Y Y U Y Y Y Y Y U Y Y Y N Y Y U U U Y Y Y

Y Y Y Y Y Y N Y Y Y Y Y Y Y Y Y U Y N Y Y N

2009 2004 1956 2008 2007 2011 2011 1982 2011 2003 1965 2008 1972 1966 2009 1997 1970 1999 1950 2009 1959 2013

a

Fish and goats. A few strains may show light pigmentation. Birds. d Sequence outside the 441-bp Telenti fragment. e Humans and swine. f Late yellow pigment g Some strains show pink pigment. h Y, yes; N, no; U, undetermined; Q, questionable; —, not grown on artificial media. i Eels. j Fish. b c

made up one-third of mycobacterial isolates reported in the 1980s (prevalence of 1.8/100,000) and that by the early 1990s they accounted for up to two-thirds of mycobacterial isolates (27–29). A more recent study correlating clinical and microbiologic data of Oregon residents who had at least one NTM isolate between 2005 and 2006, using the American Thoracic Society (ATS) and Infectious Diseases Society of America (IDSA) criteria for defining NTM infection, revealed a prevalence of 8.6 per 100,000 in the general population and 20.4 per 100,000 for individuals ≥50 years of age (30). The most common infections with NTM are pulmonary diseases, but skin and soft tissue, lymphatic, and disseminated infections also occur. Disseminated infections most often occur in the setting of advanced HIV disease, but non-HIV-infected patients can also be affected. Although regional variations in species isolation have been shown, Mycobacterium avium complex (MAC) strains are the most commonly isolated, pathogenic, slowly growing NTM, but numerous other NTM species also cause disease. In the United States, M. kansasii is the second most frequently recovered pathogenic species (24).

CLINICAL SIGNIFICANCE Because of the presence of multiple species of NTM in the environment and their opportunistic pathogenic nature, the determination of the clinical significance of the isolation of these species is based upon multiple factors, including the clinical setting, host-specific factors, the species, the

pathogenic potential of the organism, the number of positive cultures, the source of the culture isolate, and quantitation of the organisms detected by smear and culture (24). For example, although the incidence of a specific NTM such as M. gordonae in cultures is high, the pathogenicity of this species is very low, in contrast to that of species such as MAC and M. kansasii (31). In contrast to NTM, the isolation of MTBC is always an important finding in a clinical laboratory. The finding of this complex has vital epidemiologic and public health consequences. Further details on clinical significance of the MTBC may be found in chapter 30 in this Manual.

DIRECT EXAMINATION Microscopy One of the first, easiest, and least expensive means of detecting the presence of mycobacteria in clinical samples has been microscopic examination. Special acid-fast stains, along with the use of bright-field and fluorescence microscopy, are needed for staining of the organisms since the routine Gram stain is not optimal for staining mycobacteria. Further discussion of the specific techniques can be found in chapter 30 of this Manual.

Antigen Detection Antigen detection is generally not performed for direct detection of mycobacteria in diagnostic laboratories. However, more recently antigen detection assays that detect lipoarabi-

Important properties of selected slowly growing NTM speciesh

TABLE 2

Species M. avium subsp. avium, M. avium subsp. paratuberculosis, M. avium subsp. silvaticum M. intracellulare

Optimal growth temp or range (°C)

Colony morphology

Niacin

Nitrate reduction

30–37

Smooth (rough)

Neg

Neg

30–37

Smooth (rough)

Neg

Neg

Other MAC members M. arosiense

42

Smooth

ND

Pos

M. chimaera

25–37

Smooth

Neg

Neg

M. colombiense

20–37

Rough

Neg

M. vulneris

37

Smooth

Neg

Neg (85%) Neg

Other slowly growing NTMs M. bohemicum 37–40

Smooth

Neg

Neg

M. celatum

33–42

Smooth

Neg

Neg

M. genavense

31–42

Smooth

Neg

Neg

M. gordonae

30–37

Smooth/ rough

Neg

Neg

M. haemophilum

28–32

Rough

Neg

Varf

M. heidelbergense M. hiberniae

30–37 37

Smooth Rough

Neg ND

Neg Pos

M. interjectum

31–37

Smooth

Neg

Neg

Features of 16S rRNA gene (GenBank accession no. and/or reference)

Clinical relevance,a specimens of first isolation, or important laboratory feature(s)

3 subspecies established (188), another subspecies proposed (“M. avium subsp. hominissuis”) (74); identical 16S rRNA gene sequences for all subspecies (GQ153272) Several sequence variants known, some elevated to species level (see following rows) (GQ153276)

Lymphadenitis in children; pulmonary disease in adultsa; often disseminated infection in HIV patients (all caused by M. avium subsp. hominissuis) Pulmonary disease in adultsa; often disseminated infection in HIV patients

6-bp difference from M. intracellulare (EF054881) 1-bp difference (position 403) from M. intracellulare (AJ548480) 7-bp difference from M. intracellulare (AM062764) 3-bp difference from M. colombiense (EU834055)

Osteomyelitis in a child (189)d

Unique 16S rRNA gene sequence (98) (U84502) 3 sequence types are published; type 2 is very different from type 1, type 3 is similar to type 1 (194, 195); unclear taxonomic situation; possesses 2 rRNA operons (196) (L08169, type 1; L08170, type 2; Z46664, type 3) Unique 16S rRNA gene sequence (197) (X60070)

Lymphadenitis in children (193)

Several sequence variants of the 16S rRNA gene (75) (AJ581472) Sequence variants (FJ418069,e EU486080, GUI42930) Unique 16S rRNA gene sequence (X88923)

Unique 16S rRNA gene sequence (X70960) Unique 16S rRNA gene sequence (1-bp difference from M. engbaeckii) (AY438069e) Unique 16S rRNA gene sequence (X70961)

Probably similar to M. intracellulare (190)b HIV patients, blood, sputum (191)c Lymphadenitis in a child and infection after a dog bite (192)d

Pulmonary disease in adultsa

First detection in HIV patients; single cases also from nonimmunocompromised patients; almost no growth on solid media, scarce growth in liquid media, growth enhanced by Mycobactin J

Present in running water systems; usually without clinical relevance Lymphadenitis in children; skin lesion with immunosuppression; growth usually dependent on the addition of hemin to the medium Lymphadenitis in a child (198) From environmental specimens; usually without clinical relevance Lymphadenitis in a child (199)

573

(Continued on next page)

574

Important properties of selected slowly growing NTM speciesh (Continued)

TABLE 2

Species

Optimal growth temp or range (°C)

Colony morphology

Niacin

Nitrate reduction

Features of 16S rRNA gene (GenBank accession no. and/or reference)

Clinical relevance,a specimens of first isolation, or important laboratory feature(s)

Unique 16S rRNA gene sequence (X67847) 6 subspecies differing in several genes (76, 168); subspecies 1 is prevalent worldwide (171, 201, 202) (AF480601) 3 sequence variants (X80769, X80770, X93995)

Isolated from sputum specimens (200) Pulmonary disease in adultsa; rarely associated with lymphadenitis in children

M. intermedium M. kansasii

31–37 35–37

Smooth Rough

Neg Neg

Neg Pos

M. lentiflavum

22–37

Smooth

Neg

Neg

M. malmoense

30

Smooth

Neg

Neg

Unique 16S rRNA gene sequence yet very similar to that of M. szulgai (GQ153278)

M. marinum

30

Rough (smooth)

Varg

Neg

M. scrofulaceum

37

Smooth

Neg

Neg

16S rRNA gene sequence has only a 2- to 5-bp difference (none in first 500 bp) from that of M. ulcerans (203) (AJ536032) Unique 16S rRNA sequence (GQ153271)

M. shimoidei

37

Rough

Neg

Neg

M. simiae

37

Smooth

Var

Pos

M. szulgai

37

Neg

Pos

M. terrae complex (other clinically significant species include M. arupense, M. heraklionense, M. kumamotonense)

25–37

Rough/ smooth Rough/ smooth

Neg

Pos/Neg

M. ulcerans

30

Rough

Neg

Neg

M. xenopi

40–45

Smooth

Neg

Neg

Unique 16S rRNA gene sequence (no correct sequence available) Unique 16S rRNA gene sequence (GQ153280) Unique 16S rRNA gene sequence, yet very similar to that of M. malmoense (AF547969e) Group of species with similar phenotypic characteristics and clinical significance; require 16S rRNA gene sequencing for species ID

16S rRNA gene sequence has only a 2-bp difference from M. marinum (nucleotides 1248 and 1289) (203) (no correct sequence available) Unique 16S rRNA gene sequence (AJ536033 [at positions 182 and 408, C or T may be present])

a

Clinical relevance in pulmonary specimens must be proven according to the ATS guidelines. AccuProbe test is positive for MAC- and for M. intracellulare-specific probes but negative for M. avium-specific probe. AccuProbe test is positive for MAC-specific probe but negative for M. avium- and for M. intracellulare-specific probes. d No data available for AccuProbe tests. e Sequence data exist for only a fragment of the gene. f Different reports with inconsistent data. g See reference 204. The test method differed from the current strip method. h Abbreviations: Neg, negative; Pos, positive; ND, not determined; Var, varies. b c

Most isolates obtained from pulmonary specimens (77)a Lymphadenitis in children; pulmonary disease in adultsa; rare in USA but common in North European countries Swimming pool granuloma; growth rate differentials: M. ulcerans, extremely slow growing; M. marinum, 40 mycobacterial species to date. Detailed information on all genome projects either completed or still in progress can be obtained at http://www.ncbi.nlm.nih.gov/sites/en

trez?Db=genomeprj&Cmd=Search&TermToSearch= txid176.

Single Gene or 16S-23S rRNA Spacer Region Analyses Partial sequence analysis of selected genes or sequencing of the 16S-23S rRNA internal transcribed spacer region (ITS) is the most practical method for identification of mycobacterial species. Sequence variability among species, but homogeneity within species, is the basic prerequisite for this application. For several genes (16S rRNA, hsp65, and rpoB), specific regions within the gene that work well for species identification have been identified (34, 41, 59–66). The most important target for identification of slowly growing mycobacteria is the 16S rRNA gene.

16S rRNA Gene For routine identification of mycobacteria, sequence analysis of the complete 16S rRNA gene (approximately 1,500 bp) is not practical and also usually not necessary. The information content of a sequence stretch of approximately 500 bp located at the 5′ end of the gene is sufficient for identification of most species and is the only sequencing kit currently commercially available. Specific primers for amplification and sequencing of the mycobacterial 16S rRNA gene (Table 5) have been described and validated (61, 67–74). Primers 264 and 247 target genus-specific sequence regions and can be used for the detection of mycobacteria even in the presence of contaminating bacteria. Furthermore, the 285 and B9 primers can be used for the sequencing reaction. Either of these can be combined with any of the reverse

TABLE 5 Primers for amplification and sequencing of several mycobacterial target genes/sequences Target 16S rRNA 16S rRNA 16S rRNA 16S rRNA ITS 1 ITS 1 ITS 1 ITS 1 hsp65 hsp65 gyrB gyrB rpoB rpoB

Primer—orientationa 285—F 264—R B9—F 247—R Sp1—F Sp2—R ITS 1—F ITS 2—R Tbll—F Tb12—R MTUB—F MTUB—R Myco—F Myco—R

a

F, forward; R, reverse.

Sequence 5′-GAGAGTTTGATCCTGGCTCAG 5′-TGCACACAGGCCACAAGGGA 5′-CGTGCTTAACACATGCAAGTC 5′-TTTCACGAACAACGCGACAA 5′-ACC TCC TTT CTA AGG AGC ACC 5′-GAT GCT CGC AAC CAC TAT CCA 5′-GAT TGG GAC GAA GTC GTA AC 5′-AGC CTC CCA CGT CCT TCA TC 5′-ACCAACGATGGTGTGTCCAT 5′-CTTGTCGAACCGCATACCCT 5′-TCG GAC GCG TAT GCG ATA TC 5′-ACA TAC AGT TCG GAC TTG CG 5′-GGCAAGGTCACCCCGAAGGG-3′ 5′-AGCGGCTGCTGGGTGATCATC-3′

Product size (bp) ∼1,000 ∼550 ∼300

Reference 61 61 205

441

81 81 76 76 86

1,020

94

723

34

∼400

580 n BACTERIOLOGY

primers. Both forward primers are located upstream of hypervariable regions A and B, which enable the species-specific identification of most mycobacterial species (61). A few slowly growing species have identical hypervariable regions A and B or an identical complete 16S rRNA gene sequence. These include (i) M. marinum and M. ulcerans and (ii) M. kansasii (sequence variants I and IV) and M. gastri. For accurate identification of these species, sequence analysis of other genes or regions (e.g., ITS, hsp65, and rpoB) is required. All members of the MTBC have identical 16S rRNA gene sequences and thus cannot be discriminated by this technique. In addition, some species have minor intraspecies 16S rRNA gene sequence variants that differ in a few base pairs (e.g., M. gordonae, M. kansasii, and M. lentiflavum [75–77]).

techniques. However, technical difficulties, such as small size differences between the fragments or the incidence of similar or identical restriction patterns for some species of closely related mycobacteria and, in particular, new species of mycobacteria, are problems encountered with this method. With the greater availability and decreasing cost of sequencing, sequence analysis of this hsp65 gene fragment has been increasingly used instead of the PCR-restriction enzyme analysis (63, 87). Advantages of sequencing hsp65 rather than the 16S rRNA gene are especially evident in the identification of closely related species, as hsp65 is much less conserved than the 16S rRNA gene. hsp65 sequencing allows for differentiation of M. marinum from M. ulcerans, M. gastri from M. kansasii, and, with some restrictions, M. avium subsp. avium from M. avium subsp. hominissuis.

23S rRNA Gene The 23S rRNA gene is also known to contain conserved and variable sequence regions that enable the specific amplification and species identification of mycobacteria (78, 79). Variable regions can also be found in the 5′ region. The disadvantage of this target is the length of the gene (3,100 bp), which is not readily analyzed for most of the Mycobacterium species since amplification and sequencing must be performed with multiple sets of primers (79).

ITS 1 Region Another target is the internal transcribed spacer sequence, which separates the 16S and 23S rRNA genes in the operon and is denominated ITS 1. The sequence of this fragment comprises only 200 to 330 bp and thus can easily be analyzed. Several sets of primers that enable the amplification and sequencing of the complete fragment have been published (Table 5) (76, 80, 81). Primers Sp1 and Sp2 allow the genus-specific amplification of the region. Since they target the start site of the ITS, they are not optimal for analysis of the whole ITS 1 sequence. Primers ITS 1 and ITS 2 are located in the 16S and 23S rRNA gene regions, respectively. Using these primers, the entire ITS can be amplified and sequenced. However, they are not genus specific and cannot be used when cultures of mycobacteria are contaminated with other bacteria. For the ITS 1 sequence, a high variability that could be used for species identification has been shown (80, 82– 84). However, for some species, mainly for rapidly growing species but also for some slowly growing species (M. simiae and M. xenopi), two or more sequence variants have been observed. As with the 16S rRNA gene sequence, M. marinum and M. ulcerans have identical ITS 1 sequences and thus cannot be differentiated by this analysis.

hsp65 Gene One of the first genetic targets used for the differentiation of mycobacteria is an approximately 440-bp fragment of hsp65, which codes for the 65-kDa heat shock protein and is also known as the groEL2 gene (64). The amplification of this fragment, followed by a restriction enzyme digestion using the restriction enzymes BstEII and HaeIII and by analysis of the obtained digestion products using agarose gel electrophoresis, provides restriction fragment length polymorphism (RFLP) patterns that are specific for most clinically significant species (Table 5) (85, 86). An algorithm for the differentiation of the most clinically important species showing the apparent molecular sizes of the fragments has been devised. This PCR-restriction enzyme analysis technique has been used widely, since this is a simple and rapid method with no need for more-sophisticated sequencing

rpoB Gene

The rpoB gene encodes the β subunit of the bacterial RNA polymerase. For M. tuberculosis, mutations in an 81-bp region of rpoB (rifampin resistance-determining region) are known to confer resistance to rifampin. For NTM species, sequence variability within rpoB can be used for species identification (34, 60, 88). Several different sequence fragments of the approximately 3,600-bp gene have been used for amplification and sequence determination. Kim et al. (60) amplified a fragment comprising 306 bp at positions 1362 to 1668 (referred to as region 2/3), whereas Lee et al. (88) targeted a 360-bp sequence at positions 902 to 1261 (referred to as region 1/2) (34). In contrast, Adékambi et al. (34) chose a fragment more distant (region 5) and analyzed an approximately 760-bp fragment at positions 2573 to 3337 (Table 5). In the studies of Kim et al. (60) and Lee et al. (88), type strains of many slowly growing mycobacteria were included, whereas detailed analyses using region 5 (34) have been performed mainly for rapidly growing mycobacteria. Extensive investigations using clinical isolates of slowly growing mycobacteria are not available for any part of the rpoB gene to date.

Use of Sequence Databases For identification of mycobacteria by sequencing analysis, public databases are frequently used, which can present challenges. Public databases contain sequences from strains that are not validly published as species, sequences from strains identified as known species but having divergent sequences, and many sequences from uncultured bacteria that are not further characterized. Furthermore, old sequences that are fragmentary or faulty are still included in the public databases (89). Thus, the evaluation of the search result has to be done carefully. For example, in the case of an uncommon species, cross-checking of the valid description as a species can easily be performed by using the List of Prokaryotic Names with Standing in Nomenclature (http://www.bacterio.net). When obtaining several correct results deriving from different species, it may help to check the database entries if the sequences have been obtained from type strains. The International Nucleotide Sequence Database Collaboration database (INSDC; http://www.insdc.org/) can be used for the analysis of the sequences derived from the different genes as detailed above. There are some curated databases and analysis tools that have been constructed for the analysis of the 16S rRNA gene sequences, such as the commercially available tools SmartGene IDNS (SmartGene, Lausanne, Switzerland), RipSeq (Isentio, Sunnyvale, CA), and the MicroSeq microbial identification system (Applied Biosystems, Foster City, CA) or two tools with free access, the Ribosomal

31. Characteristics of Slowly Growing Mycobacteria n 581

Database Project (RDP; http://rdp.cme.msu.edu/) and the RIDOM (Ribosomal Differentiation of Medical Microorganisms) database (http://www3.ridom.de/rdna/). The advantage of these databases is the quality control of the entries, ensuring that most of the difficulties associated with public databases will not be encountered. For the commercial Webbased databases, the user also has the option to update the system as needed. However, these curated databases can lag behind the updates of the INSDC database and may not include sequence variants of species.

Pyrosequencing Pyrosequencing differs from Sanger sequencing in that it relies on the detection of pyrophosphate release on nucleotide incorporation rather than chain termination with dideoxynucleotides. Identification of mycobacteria by pyrosequencing relies on a 20- to 30-bp region within the hypervariable region A of the 16S rRNA gene (90–92). A study evaluating pyrosequencing for the analysis of AFB isolates was performed by Bao et al. in comparison to phenotypic analysis and Sanger sequencing. Pyrosequencing was capable of identifying 114 (98%) of the 117 AFB isolates and correctly identified most of the slowly growing mycobacteria to the species level (92). Of the mycobacteria tested, it was not able to identify M. scrofulaceum and M. simiae to the species level. Similar to Sanger sequencing of the 16S rRNA gene, pyrosequencing has the same limitation for identification of some of the closely related organisms, as discussed in “Single Gene or 16S-23S rRNA Spacer Region Analyses” above. Pyrosequencing has the advantage of being a rapid, simple, and inexpensive molecular method for identification of Mycobacterium species. Additional sequencing may be required to differentiate some species due to the short sequence length generated by this technique (91, 93).

GENOTYPIC IDENTIFICATION OF SPECIES OF THE MTBC Within the MTBC, the most prevalent species is M. tuberculosis (∼93 to 97%), followed in frequency by M. bovis and M. bovis BCG. Identification of M. bovis is of clinical relevance because of the inherent resistance of this species to pyrazinamide, one of the first-line agents for treatment of TB. Strains that are monoresistant to pyrazinamide should be analyzed by molecular techniques. These strains can be M. bovis, M. bovis BCG, or, although rare, monoresistant M. tuberculosis strains. Moreover, M. bovis BCG is used for treatment of bladder cancer, and thus correct identification of M. bovis BCG isolated from patients treated for bladder cancer is necessary to enable a correct decision for treatment of a patient. The identification of the other species is mainly of epidemiological importance and may be an indication of the source of the infection.

gyrB Gene Sequencing The gyrB gene encodes the B subunit of DNA gyrase (topoisomerase II), an enzyme essential for bacterial replication. In 2000, Kasai et al. (94) showed single nucleotide polymorphisms (SNPs) in an approximately 1.2-kbp fragment of the gyrB gene which were specific for some species of the MTBC (Table 5). More-detailed analyses confirmed these results and determined that they could be extended to most members of the MTBC (95).

RD/Spoligotyping Although all members of the MTBC are characterized by a high genetic similarity, several sequence polymorphisms

that can be used for species identification have been discovered. Comparative genomic analysis has shown that members of the MTBC have evolved from a common ancestor through sequential DNA deletions with precise genomic deletions (96). These areas in the genome are exploited for the identification of the members of the MTBC and are referred to as regions of difference (RD). The presence or absence of these deletions can be analyzed by PCR assays (97). Spoligotyping, a technique applied primarily for strain typing, identifies most species within the MTBC. However, with this technique, M. bovis BCG cannot be discriminated from M. bovis.

Line Probes Specific for MTBC A commercially available assay (GenoType MTBC; Hain Lifescience, Nehren, Germany) based on line probe technology enables the identification of species within the MTBC (98–101). The test can be performed from solid or liquid media, with a total test time of 4 to 6 hours. The assay is based on an MTBC-specific 23S rRNA gene fragment, gyrB DNA sequence polymorphisms, and the RD1 deletion of M. bovis BCG. Specific oligonucleotides targeting these polymorphisms are immobilized on membrane strips. Amplicons derived from a multiplex PCR react with these probes during hybridization. Species can be identified according to the interpretation table provided with the kit. The inclusion of an MTBC-specific 23S rRNA gene fragment confirms the presence of MTBC in order to rule out possible cross-reactivity. Specific patterns can be obtained for M. tuberculosis/M. canettii, M. africanum/M. pinnipedii, M. microti, M. caprae, M. bovis, and M. bovis BCG. M. tuberculosis and M. canettii as well as M. africanum and M. pinnipedii share identical gyrB sequences, and thus they can be identified by specific patterns, but they cannot be differentiated from each other. The specificity of this test was 100% when strains from culture collections or clinical isolates were compared (99– 101). With strains from culture collections, all species, with the exception of an M. canettii strain (identified as M. tuberculosis), were correctly identified (101). Additionally, 100% of the clinical strains studied were correctly identified (99–101). No false-positive results were obtained with several NTM strains (101).

Real-Time PCR Assays Laboratory-developed, multiplex real-time PCR assays have been described for the rapid identification of MTBC members to the species level from both culture and clinical specimens. The presence or absence of regions of difference (RD) allows identification to the species level of the MTBC using real-time PCR analysis (96, 102).

Other Molecular Tests for NTM and MTBC Sequence analysis of target genes for the identification of mycobacteria may not be practical for routine clinical laboratories. Commercially available assays that are based on liquid- or solid-phase hybridization have been shown to be easily implemented into a routine workflow. They are intended for the detection of some of the most important Mycobacterium species and can be performed from both solid and liquid media.

AccuProbe Culture Identification Tests The AccuProbe test was the first commercial molecular assay for the identification of selected mycobacterial species from positive culture media. Probes are available for the identification of MTBC, M. avium, M. intracellulare, MAC,

582 n

BACTERIOLOGY

M. gordonae, and M. kansasii. All probes are FDA cleared and commercially available. Species identification is based on the hybridization of specific DNA probes to rRNA of the bacteria. Briefly, by heat treatment and sonication, nucleic acids, including the target 16S rRNA, are released from the mycobacteria. A specific DNA probe hybridizes with the target rRNA. Finally, the DNA-rRNA hybrid molecule can be detected by chemiluminescence. The results are obtained within 2 hours. The utility of these tests has been proven in many studies and by usage in many laboratories worldwide (76, 103–107). Specificities of the tests are usually reported to be 100%, or sometimes lower (96%), for MAC. Sensitivity values vary with the test used: 100% for M. gordonae, 95.2% for MAC, and 97.4 to 100% for M. kansasii (76, 103, 104, 107). It has also been shown that MTBC organisms can reliably be identified in the presence of M. avium (105). However, there are also some disadvantages of these tests. The probes are limited to a few, although important, mycobacterial species, necessitating the performance of additional tests for identification of species for which there are no probes. The need to perform individual tests for each target species renders the tests expensive and, if not performed in parallel, also time-consuming. False-positive results for MAC probes have been reported, but they can be prevented if a higher cutoff value (80,000 relative light units instead of the 30,000 relative light units recommended in the technical insert) is used (108). Studies have also shown cross-reactions with the M. intracellulare probe and several slowly growing mycobacterial species (M. arosiense, M. chimaera, M. nebraskense, and M. saskatchewanense) (109). Similarly, cross-hybridizations have been documented with the MAC probe and M. nebraskense, M. palustre, M. saskatchewanense, and M. paraffinicum. Cross-reactivity has been seen with the MTBC probe and M. celatum (110) and M. holsaticum (109), although this cross-reactivity can be avoided when the selection step of the procedure is performed for 10 min, according to the manufacturer’s instructions (111).

Line Probe Assays Alternatively, techniques based on the application of PCR plus reverse-hybridization-designed DNA strip assays (line probe assays), have been developed. Briefly, the target sequences are amplified by PCR using biotinylated primers. The amplified PCR products are allowed to hybridize to immobilized, membrane-bound probes covering the speciesspecific sequence fragment, followed by an enzyme-mediated color reaction. The banding patterns can be analyzed by eye by comparing the patterns on an interpretation chart. The identification of the species relies on specific banding patterns (Fig. 5). Three tests for the identification of several mycobacterial species are commercially available but are not yet FDA cleared. The INNO-LiPA Mycobacteria v2 assay (Innogenetics, Ghent, Belgium) is based on the nucleotide differences in the 16S-23S rRNA gene spacer region and can detect the following slowly growing Mycobacterium species: MTBC, M. kansasii, M. xenopi, M. gordonae, M. genavense, M. simiae, M. marinum, M. ulcerans, M. celatum, M. avium/ M. intracellulare/M. scrofulaceum complex, M. avium, M. intracellulare, M. scrofulaceum, M. malmoense, and M. haemophilum. The GenoType Mycobacterium CM/AS test (Hain Lifescience, Nehren, Germany), is based on the detection of species-specific sequences in the 23S rRNA gene. It is composed of two different strips (strip CM is for common mycobacteria, and strip AS is for additional species) and can identify the following slowly growing mycobacteria: with

FIGURE 5 Line probe assay (GenoType Mycobacterium CM/AS) for the identification of Mycobacterium species. CC, conjugate control; UC, universal control; GC, genus-specific control; 4 to 17, specific bands for identification of the species; M, marker line (flags the upper side of the strip). Lines 1 to 6, CM strips; lines 7 and 8, AS strips. First lane, M. avium; second lane, MTBC; third lane, M. gordonae; fourth lane, M. malmoense; fifth lane, M. xenopi; sixth lane, negative control; seventh lane, M. genavense; eighth lane, M. haemophilum. doi:10.1128/9781555817381.ch31.f5

strip CM, M. avium, M. gordonae, M. intracellulare/M. chimaera, M. scrofulaceum/M. paraffinicum/M. parascrofulaceum, M. interjectum, M. kansasii, M. malmoense/M. haemophilum/M. palustre/M. nebraskense, M. marinum/M. ulcerans, MTBC, and M. xenopi; and with strip AS, M. simiae, M. celatum (types 1 and 3), M. genavense/M. triplex, M. lentiflavum, M. heckeshornense, M. szulgai/M. intermedium, M. haemophilum/M. nebraskense, M. kansasii, M. ulcerans, M. gastri, M. asiaticum, and M. shimoidei (a slash separating two or more species means that they share the same pattern). Both the Innogenetics and Hain Lifesciences tests can be conducted on isolates from liquid or solid culture media, and the results are available in approximately 4 to 6 hours. Several studies have evaluated the usefulness of the tests. Concordant results of 89.9 to 98.6% for the GenoType CM and AS assays with standard methods (e.g., sequencing) have been reported (112–117). Most (up to 96%) of the regularly encountered species already can be identified using the GenoType CM assay alone (116). A limitation of this test is the resolution of patterns that are shared by more than one species, which necessitates additional techniques for further discrimination (e.g., sequencing of additional gene targets or biochemical tests) leading to a delay in obtaining a final result. For the INNO-LiPA assay, 92.2% to 98.4% concordant results with standard methods have been reported (115, 118). The assay can further differentiate M. kansasii and M. chelonae strains into subtypes. The clinical significance of this information so far is unclear, and thus this knowledge is mainly of epidemiological value. Using line probe assays, more than 90% of the commonly encountered mycobacteria can rapidly be identified. In a few cases, false identification has been reported (109, 116, 118). For the GenoType AS assay, results for M. celatum should be reported only if they are obtained from solid cultures with appropriate growth rates and typical colony morphology. Recent studies have shown that the GenoType

31. Characteristics of Slowly Growing Mycobacteria n 583

M. intracellulare probe cross-hybridizes with several slowly growing mycobacterial species, including M. arosiense, M. chimaera, M. colombiense, M. mantenii, and M. saskatchewanense. Two infrequently encountered NTM species, M. riyadhense and the nonvalidated species “M. simulans,” were incorrectly assigned to the MTBC by the GenoType assay (109). Similarly, the INNO LiPA MAIS assay cross-reacts with M. arosiense, M. heidelbergense, M. mantenii, M. nebraskense, M. parascrofulaceum, and M. paraffinicum (109). Importantly, neither of the line probe systems has been reported to misidentify members of the MTBC as an NTM (109). All results obtained with line probe tests should be confirmed by key established phenotypic characteristics, such as growth rate, colony morphology, and pigmentation. A third line probe assay was recently described, the Speed-oligo Mycobacteria (Vircel, Granada, Spain), and is based on targeting both 16S rRNA and 16S-23S rRNA regions. The assay can identify 19 mycobacterial species, including MTBC (M. tuberculosis, M. bovis, M. microti, M. africanum, M. caprae) and 10 additional NTMs (M. avium, M. gordonae, M. interjectum, M. intracellulare, M. kansasii, M. malmoense, M. marinum/M. ulcerans, M. scrofulaceum, and M. xenopi). The Speed-oligo Mycobacteria system has a limited spectrum of species identification compared to the other assays and identifies many of the mycobacterial species to the complex or group. It can be performed on isolates from solid or liquid media within 3 hours. The only clinically relevant misidentification reported in the limited studies published to date was the misidentification of M. marinum as M. kansasii (119, 120). The Speed-oligo Mycobacteria system has also been evaluated for use directly from respiratory specimens and demonstrated an overall sensitivity of 76% (93% from smear-positive and 56% from smear-negative specimens) and specificity of 99%. The study concluded that the assay is an easy and fast alternative for detecting mycobacteria and differentiating between MTBC and NTM, especially from smear-positive specimens (121).

Multiplex Real-Time PCR Assays There are a handful of reports in the literature on the use of multiplex real-time PCR with probe-based detection for the rapid identification of MTBC and NTMs from culture isolates (122) or for the differentiation of members of the MTBC (123, 124). More recently, multiplex real-time PCR coupled with high-resolution melting curve analysis has been reported for the detection of mutations in MTBC isolates associated with resistance to first- and second-line antituberculous agents (125, 126). A commercial multiplex real-time PCR assay (Seegene, Seoul, South Korea) uses a proprietary technology that enables simultaneous screening and detection of multiple targets using a single fluorescent channel. This technology has been used for discrimination between NTM and MTBC directly from clinical specimens using the AnyplexTM MTB/NTM assay. In addition, the Seegene Anyplex II MTB/MDR/XDR detects MTB, MDRTB, and extensively drug-resistant (XDR) TB in a single real-time PCR from clinical specimens. Although the Seegene assays are not U.S. FDA cleared, they have received CE certification. Studies demonstrating the clinical utility of the Seegene multiple PCR assays for detection of Mycobacterium species are yet to be performed.

M. leprae PCR As M. leprae is a nonculturable organism, PCR techniques have been developed and applied for its detection from a variety of specimen sources, such as slit-skin smears, blood, nasal cavity, biopsy specimens, and urine (127–132). Molec-

ular targets for detection of M. leprae PCR have included protein antigens of 18 kDa, 36 kDa (known as the prolinerich antigen [pra]), and 65 and 85 kDa (the 16S rRNA gene and repetitive sequences) (127, 128, 130, 132–136). Many studies have shown that PCR methods are specific and more sensitive than conventional methods (split skin smears, histopathology, and serologies) for the diagnosis of M. leprae (127, 128, 130, 132–137).

Mass Spectrometry Matrix-assisted laser desorption ionization–time of flight mass spectrometry (MALDI-TOF MS) is revolutionizing the identification of microorganisms and has been recently applied to the identification of mycobacteria. MALDI-TOF MS analysis of mycobacteria involves several steps including inactivation, extraction, and analysis. Several different methods have been described for the inactivation and extraction of mycobacterial cells. Since there are additional safety concerns when dealing with acid-fast bacillus (AFB)positive cultures, these methods differ from the extraction methods described for other bacteria and fungi. All inactivation and extraction steps should be performed within a BSL3 facility using the appropriate personal protective equipment (PPE). MALDI-TOF analyses of both whole-cell (138) and purified protein extraction (139–141) methods have been described. Inactivation of mycobacterial cells is usually achieved either by ethanol or by heat treatment and dissociation/lysis of cells by mechanical lysis using either a micropestle or bead beating techniques. Extraction is performed by adding formic acid and acetonitrile to the inactivated mycobacterial cells (139, 141). Following inactivation and extraction, an aliquot is spotted onto a solid target support plate and overlaid with a chemical matrix (for example, alpha-cyano-4-hydroxy-cinnamic acid). The sample plate is loaded into the instrument, and mycobacterial proteins are ionized using a laser. The ionized proteins then move through the time of flight chamber and are separated based on the mass-to-charge ratio of the ions, such that the lighter proteins hit the detector first, followed by the heavier proteins. A mass spectrum is produced as the ions collide with the ion detector and serves as a “fingerprint” for identification of the mycobacterial species. Currently, there are two commercial MALDI-TOF platforms available for microbial identification, the Vitek MS (bioMerieux, Durham, NC) and the Biotyper system (Bruker Daltonics, Billerica, MA) (138–141). Databases supplied with both instruments contain entries for MTBC and many NTMs, including those most commonly encountered in the clinical laboratory. MALDI-TOF MS databases continue to improve in terms of species coverage and the depth of strain diversity, but more esoteric species may be underrepresented and may need to be added by the end user or may await future database updates by the manufacturers. Recent studies have demonstrated the ability of MALDITOF MS to serve as a rapid and accurate means for the identification of mycobacteria from both solid and liquid media (138–141). Saleeb et al. (142) reported some of the earliest use of MALDI-TOF MS with the identification of 104 mycobacterial isolates comprising 17 species. Since this early report, others have also demonstrated that MALDITOF MS can unambiguously identify the most commonly encountered species of mycobacteria (143). However, species of the MTBC cannot be differentiated and the closely related M. chimaera and M. intracellulare are not separated, according to recent studies (138, 144). A potential pitfall of the identification by MALDI-TOF MS in comparison to molecular sequencing methods is the

584 n

BACTERIOLOGY

requirement for a moderate amount of growth present on solid media rather than the scant growth required for sequencing. Not surprisingly, this problem is most evident when dealing with the slowly growing mycobacteria (141). In such circumstances, sequencing may provide a more rapid turnaround time (TAT; i.e., 12 to 24 h) for identification than MALDI-TOF analysis. Although the costs of MALDI-TOF MS instrumentation are not insignificant, studies have shown that this technique has minimal associated reagent costs, requires minimal technologist time, can be utilized for a wide variety of microorganisms, and provides a reduction in TAT for organism identification, leading to an overall reduction in costs (145). A second MS method is a novel technology coupling PCR amplification and electrospray ionization mass spectrometry (ESI-MS) to identify M. tuberculosis complex, MTBC resistance determinants, and NTM from culture within ∼6 hours. This system measures the mass/charge ratio of PCR amplicons generated from several loci, focusing on conserved, species-specific regions and resistance determinants, to identify base compositions comparative to a database of mycobacteria and resistance determinants. Using the base composition as unique molecular signatures (“fingerprints”), the PLEX-ID system (Abbott Molecular, Des Plaines, IL) is able to identify the organism as an MTBC or NTM and provide resistance markers if an MTBC is identified (146, 147). A recent study demonstrated the ability of the MDR-TB assay on the PLEX-ID to appropriately identify all MTBC and all slowly growing NTMs tested from both broth (MGIT tubes) and solid media using database version NFDU.415.455.349. In comparison to the agar proportion method, the sensitivity and specificity for the detection of MTBC drug resistance using the MDR-TB assay were 100% and 92.3% for rifampin, 100% and 93.8% for isoniazid, 91.0% and 94.4% for ethambutol, and 100% and 100% for fluoroquinolones, respectively (148). Further studies need to be performed to determine if the MDR-TB assay is capable of detecting AFB directly from clinical specimens. High instrument costs of the PCR ESI-MS system may hamper the implementation of this technology into small clinical microbiology laboratories, and therefore this technology may be limited to use in reference laboratories or state public health laboratories.

TYPING SYSTEMS Historically, the first strain typing method for M. avium was serotyping based on seroagglutination procedures. Combined use of serotyping and species-specific DNA probes has shown that the organisms previously named serovars 1 through 6 and 8 through 11 are M. avium and that serovars 7, 12 through 17, 19, 20, and 25 are M. intracellulare. Finally, multilocus enzyme electrophoresis has been shown to provide a wider range of polymorphisms than serotyping (149). Serotyping, for both MAC and other mycobacteria, has been replaced by molecular strain typing.

Genotyping of M. tuberculosis Strains The genotyping of M. tuberculosis isolates contributes to the knowledge and control of TB by indication of epidemiological links between patients, discrimination between exogenous reinfections and endogenous reactivations, outbreak detection, and the recognition of laboratory cross-contaminations. The CDC has initiated a laboratory program to provide genotyping services for TB control programs to public health laboratories. Further information may be obtained from their website at http://www.cdc.gov/tb/prog rams/default.htm.

The introduction of the IS6110-RFLP technique marked a milestone in determining the molecular epidemiology of M. tuberculosis and has been the reference standard for many years. More recently, several alternative PCR-based techniques, which are less time-consuming and in part more discriminative than the classical IS6110-based technique, have been developed.

IS6110 RFLP Typing IS6110 is an insertion sequence that is present in variable numbers (0 to >20 [150]) and has been inserted at various positions in the genomes of MTBC isolates. IS6110-based RFLP analysis is performed using an internationally standardized protocol that facilitates comparison of patterns generated by different laboratories (151). High-quality DNA is isolated from mature culture isolates and subjected to restriction enzyme digestion. The DNA fragments are electrophoretically separated, transferred to a membrane, and hybridized with an IS6110 probe. The resulting patterns are compared to an internal size standard. For large-scale comparisons, the patterns should be digitalized by scanning in order for them to be stored in a database. Using computer analysis, the results from separate runs can be compared to detect identical patterns. The IS6110 fingerprint patterns are highly discriminatory, but only for patterns with seven or more bands. The complete analysis is laborious and time-consuming if the time to obtain sufficient growth on solid media and the multiple technical steps are considered. Furthermore, the analysis of the complex banding patterns requires sophisticated pattern-matching computer software.

Spoligotyping Spoligotyping (an abbreviation for spacer oligonucleotide typing) is a PCR-based technique that targets the variability in the direct repeat (DR) locus of M. tuberculosis. The DR region that is present in all MTBC strains consists of multiple DRs of a conserved 36-bp sequence separated by nonrepetitive unique spacer sequences. Of these, 43 spacer sequences identified in the genomes of M. tuberculosis and M. bovis BCG are used for the spoligotyping assay (152). The spacer segments are amplified by PCR using primers that target the DR sequence. Amplified DNA fragments are hybridized to a membrane with 43 covalently bound oligonucleotides complementary to the 43 spacers. There are several advantages of spoligotyping compared to IS6110 RFLP analysis. Since the technique is PCR based, a small amount of DNA is sufficient for analysis. Thus, spoligotyping can be performed from scantily grown solid as well as liquid cultures, enabling a more rapid turnaround time. The technique is less laborious and more rapid than the classical IS6110 RFLP method. Additionally, the format of the result can easily be transferred into a binary code that can be handled in common computer programs, rendering large-scale comparisons less difficult than the classical method. Unfortunately, spoligotyping is less discriminatory than IS6110 RFLP typing and not sufficient for populationbased studies. It can be used for M. tuberculosis strains with few IS6110 copies or for M. bovis isolates that also have few IS6110 copies. This technique is useful for the analysis of assumed laboratory cross-contaminations, as the results can be obtained rapidly.

MIRU-VNTR Typing Genotyping may be performed based on the variable numbers of tandem repeats (VNTR) of different classes of genetic elements named mycobacterial interspersed repetitive units

31. Characteristics of Slowly Growing Mycobacteria n 585

(MIRU) (153, 154). This technique is based on the amplification of multiple repeat loci and the subsequent determination of the amplicon size, which depends directly on the number of VNTR copies. Initially, 12 different loci were shown to be suitable for analysis and were considered to possess discriminatory power comparable to that of IS6110 RFLP analysis (155). This method was adopted in the previously discussed CDC TB genotyping program. Subsequent studies, however, have found the 12-locus assay, even when combined with spoligotyping, to be less discriminatory than the classic RFLP method. The use of a new set of 24 loci for epidemiological studies has been proposed (153, 156, 157). A subset of 15 highly discriminatory loci among the 24 loci provides sufficient discriminatory information for routine epidemiological discrimination. The amplification of the various loci is usually performed in a multiplex format. The PCR products can be analyzed using a DNA sequencing instrument. For small-scale comparisons, analysis of PCR products using classical agarose gel electrophoresis is also possible. The result format is a series of 12, 15, or 24 digits (depending on the MIRU format employed). The number designates the number of repeats at each locus. This format is portable between laboratories and thus simplifies the comparison of large databases. The MIRU-VNTR technique is rapid, with minimal growth required, and is increasingly replacing the IS6110 RFLP reference standard method.

to clinical and epidemiologic studies of M. kansasii, M. szulgai, and MAC (166–171). Recently, PFGE has been applied to study macrolide-susceptible and -resistant isolates and multiple cultures of patients with nodular MAC disease and bronchiectasis (166).

Multilocus Sequence Typing Another approach for NTM typing is analysis of multiple genes with a high sequence variation, like hsp65 or ITS. However, available sequences have not provided the level of variability needed for epidemiological analyses. With the progress of whole-genome analysis and the possibility of large-sequence determination, however, multilocus sequence typing is likely to prove increasingly useful for typing of NTM.

Repetitive-Unit Sequence-Based PCR Repetitive-unit sequence-based PCR (rep-PCR) has been used for outbreak control of several slowly growing mycobacterial species, including M. simiae, M. gordonae, M. terrae, M. tuberculosis, and M. avium complex (172–174). The method is commercially available through the DiversiLab System (bioMérieux, Durham, NC). Although the technology is proprietary, the company offers a Web-based library of rep-PCR sequences that can be searched for matches. Few studies have compared rep-PCR results to those obtained with PFGE.

Multilocus Sequence Typing Variable-Number Tandem Repeat Multilocus sequence typing (MLST) directly measures the DNA sequence variations in a set of housekeeping genes in M. tuberculosis and characterizes strains by their unique allelic profiles. MLST involves PCR amplification of 7 housekeeping genes (gyrA, gyrB, katG, purA, recA, rpoB, and sodA) followed by DNA sequencing (158). A recent study comparing MLST to MIRU-VNTR and spoligotyping was performed to evaluate the discriminatory power between the three methods to assess the genetic diversity of M. tuberculosis. Of the three methods, MIRU-VNTR showed the highest discriminatory power followed by spoligotyping followed by MLST. The authors of the study conclude that the combination of MIRU-VNTR and spoligotyping provides the greatest level of discriminatory power and therefore is the most useful genotyping tool to be applied to M. tuberculosis isolates (158).

Genotyping of Slowly Growing NTM Different approaches have been used for strain comparisons of NTM. Whole-genome analyses have included pulsedfield gel electrophoresis (PFGE) (159–163), amplified fragment length polymorphism analysis (AFLPA) (164), or RFLP typing using specific molecular markers (162, 165). For PFGE and AFLPA, no genetic information is necessary and the procedures can be performed without further knowledge of the species genome. For RFLP typing, some genetic information, such as the presence of specific IS elements, is a prerequisite.

Pulsed-Field Gel Electrophoresis PFGE has been a useful technique for strain typing of the slowly growing NTM. This technique requires an actively growing culture and 3 to 4 weeks for completion. Problems with standardization may result from cell clumping of rough strains. This clumping may also result in different amounts of DNA even from different batches of the same strain, causing problematic uneven (light and overloaded) lanes. Previous studies have detailed the method of PFGE related

MIRU-VNTRs were initially applied to typing of M. avium subsp. paratuberculosis (MAP) strains but now have been expanded to other members of the MAC (175–177). Although the VNTR loci are usually species specific, the low cost, reproducibility, smaller volume of DNA requirement, and speed of the VNTR technique suggest that it could be a substitute method for PFGE strain comparison with MAC (178).

IS1245/IS900 The most extensive investigations for typing of NTM strains have been undertaken for M. avium. RFLP analysis based on the insertion sequence IS1245, which is restricted to subspecies of M. avium, has been used for genotyping. Standardization of this technique was proposed in 1998 (179), but this technology never became widespread. However, Mijs et al (180) used this method as one tool for distinguishing M. avium subsp. avium strains obtained from bird specimens from swine- and human-derived strains, for which they proposed the name “M. avium subsp. hominissuis.” RFLP analysis based on the insertion sequence IS900 can be used for genotyping of MAP. The availability of the complete genome sequences of the “M. avium subsp. hominissuis” strain 104 and the MAP strain K-10 allowed the search for divergent regions that can be used for standardization of other techniques for genotyping (165). Based on the genomic sequence data, largesequence polymorphisms, which were analyzed for their variability among different M. avium strains, could be identified. By the use of multilocus sequence typing, many genes that contain multiple sequence variations can now be analyzed in parallel (181).

ANTIMICROBIAL SUSCEPTIBILITY TESTING Antimicrobial susceptibility testing (AST) of clinically relevant mycobacteria is crucial for the control of infections

586 n BACTERIOLOGY

caused by many species of mycobacteria and should be performed for all species, for which CLSI guidelines are available (182). Procedures are technically demanding and require rigid safety precautions, especially when liquid cultures are involved. When handling positive cultures with a high concentration of bacilli, cross-contamination is another critical issue. Therefore, antimicrobial susceptibility testing should be performed only by experienced personnel.

Nontuberculous Slowly Growing Mycobacteria AST of the NTM requires skill and knowledge of the individual species characteristics. Therefore, the CLSI has advised laboratories that infrequently encounter NTM to refer those isolates to a qualified reference laboratory. Laboratories that elect to perform in-house NTM AST should carefully validate their results and monitor their proficiency regularly, as required by specific accrediting agencies, such as the College of American Pathologists (182). Generally, broth micro- or macrodilution techniques in serial 2-fold concentrations are recommended. Testing is most efficiently performed in 96-well microtiter panels. Breakpoints for some species of slowly growing mycobacteria have been proposed for the following antimicrobials: rifampin, rifabutin, amikacin, ethambutol, ciprofloxacin, moxifloxacin, minocycline, doxycycline, clarithromycin, trimethoprim-sulfamethoxazole, and linezolid (182). The CLSI recommendation specifies testing for slowly growing species, including M. kansasii, MAC, and M. marinum. The CLSI and the statement of the American Thoracic Society and the Infectious Diseases Society of America have recommended that only rifampin and clarithromycin results be reported for M. kansasii unless the isolate is rifampin resistant; if so, all of the previously listed antimicrobials should be tested (24, 182). For isolates of M. marinum, experts in mycobacterial susceptibility testing concur that AST is unnecessary in most cases due to the narrow range of MICs for this species. In cases of intolerance to agents typically effective for the treatment of infections due to M. marinum (i.e., clarithromycin, rifampin, and ethambutol), antimicrobial testing of the previously listed agents should be performed (182). For MAC, because previous studies have shown that no correlation exists between MICs and the clinical response of the patient to agents other than clarithromycin, testing of only clarithromycin is recommended. However, a recent study of patients with MAC has shown a correlation of amikacin MICs to clinical response and amikacin breakpoints have been proposed to the CLSI (183). Interpretive criteria are also provided for moxifloxacin and linezolid, but no clinical correlation studies have been performed to date (182). Other species of slowly growing NTM have insufficient data to make specific recommendations for testing, and therefore the CLSI recommends that testing of these NTM should follow the guidelines for testing rifampin-resistant M. kansasii. Fastidious species that require iron- or hemin-containing media for growth, such as M. haemophilum, may be tested using an alternative agar disk elution method with commercial antimicrobial disks in molten agar. The CLSI has recommended testing of antimicrobials, including rifampin, clarithromycin, amikacin, ciprofloxacin, trimethoprim-sulfamethoxazole, linezolid, and doxycycline or minocycline (182). Specific details and discussion of the AST procedures can be found in chapter 76 of this Manual.

M. tuberculosis Complex Currently, methods for testing MTBC are based on the method of proportion, which relies on a clinical definition of drug resistance. Susceptibility testing of MTBC may be performed in agar-based (agar proportion method) or brothbased (commercial) systems. Agar proportion methods may be performed using either commercially prepared or inhouse media. Both the agar proportion and newer liquid detection systems define resistance as growth of >1% of an inoculum of bacterial cells in the presence of a “critical” concentration of the drug. By convention, the critical concentration corresponds to the lowest concentration of drug that inhibits 95% of “wild-type strains” of MTBC that have never been exposed to the antimicrobials without simultaneously inhibiting strains of MTBC from patients who do not respond to treatment and that are considered resistant. The critical concentration is thus the standard concentration by which susceptibility and resistance are established. A newer method for MTBC susceptibility testing uses a 96-well microtiter panel containing 12 lyophilized first- and second-line antimycobacterial agents. In contrast to the more common agar proportion methods, the MycoTB panel (Thermo Scientific/TREK Diagnostics, Oakwood Village, OH) tests a range of concentration of each antimycobacterial providing a MIC that is consistent with AST formats for other bacterial organisms. Two studies comparing the Sensititre MycoTB panel have demonstrated 94 to 100% categorical agreement with the reference agar proportion method. The Sensititre MycoTB panel appears to be a more rapid (10 to 14 days versus 21 days), quantitative, and efficient method than the gold standard agar proportion method (184, 185). An additional advantage to the Sensititre MycoTB panel is the availability of second-line susceptibility results in the case of a resistant strain. The disadvantage is the absence of pyrazinamide in the panel, which would require testing by another method. Another disadvantage is potential safety concerns associated with working in microtiter plates with possible drug-resistant strains. Generally, the first isolate of MTBC cultures from each patient should be tested, and susceptibility testing should be repeated if the patient fails to respond to therapy or if the cultures remain positive at 2 to 3 months. Primary susceptibility testing includes a battery of antimicrobials, including isoniazid at two concentrations (critical and higher concentrations), rifampin, ethambutol, and pyrazinamide. When an isolate is resistant to rifampin or any two of the other primary drugs, a secondary panel including a higher concentration of streptomycin and additionally capreomycin, ethionamide, amikacin, p-aminosalicylic acid, rifabutin, cycloserine, linezolid, moxifloxacin, and levofloxacin should be tested (182). Direct susceptibility testing of smear-positive samples from patients known to have or suspected of having M. tuberculosis involves inoculation of drug-containing medium with a directly processed sample (i.e., concentrated after decontamination and digestion). Indirect susceptibility testing may be performed using cultures already growing either in liquid or in solid medium. This method is usually used on smear-negative samples or if the direct test results are invalid due to contamination, insufficient numbers of colonies in the drug-free quadrants, or insufficient growth after 3 weeks of incubation. Details on both methods of susceptibility testing can be found in chapter 76. Standardized agar proportion methods are not considered rapid methods. Therefore, the addition of a commercial liquid susceptibility system with shorter incubation is strongly recommended for

31. Characteristics of Slowly Growing Mycobacteria n 587

patient testing. It is imperative, however, that any commercial system be validated to produce results that correlate with the standard susceptibility agar proportion methods. As with any other laboratory test, adequate and consistent quality control and proficiency testing should be performed to ensure accurate and consistent results (182). Molecular detection of drug resistance markers in MTBC generally makes use of laboratory-developed PCR or sequencing methods that target well-characterized genes (katG, rpoB, pncA, gyrB, etc.) that have mutations associated with resistance to first- and second-line agents (186). These LDTs have been used for detection of resistance markers from culture isolates and, in some instances, directly from respiratory specimens. In addition, the commercially available Xpert TB/RIF assay (Cepheid), discussed earlier in this chapter, provides information about rifampin resistance directly from respiratory specimens. The CDC offers a service for the rapid molecular detection of drug resistance in MTBC using sequencing methods from culture isolates or NAA test-positive sediments. Targets associated with resistance to both first- and second-line agents are interrogated with reported sensitivities of 97.1% for rpoB, 85.4% for katG, 84.6% for pncA, 78.6% for embB, 81.6% for gyrA, and 90.8% for MDR TB (187).

EVALUATION, INTERPRETATION, AND REPORTING OF RESULTS Implementation of sensitive and rapid methods for the detection, identification, and susceptibility testing of MTBC and NTM is of paramount importance to the diagnosis and control of TB and NTM disease. The use of the fluorochrome stain for mycobacterial smears, a broth-based culture system, and the use of DNA probes or sequence analysis for identification along with direct susceptibility testing of smear-positive specimens are optimal for supporting the critical decisions required for optimal patient care. As emphasized in this chapter, phenotypic testing of the slowly growing mycobacterial species has limited use and for other mycobacteria (i.e., rapidly growing mycobacteria) should be applied only in conjunction with molecular assays. Laboratory work should be organized to prioritize timely and accurate identification of MTBC and susceptibility testing of clinically significant isolates. In order to optimize rapid detection, identification, and AST for the shortest turnaround times, the U.S. Association of Public Health Laboratories has recommended that the Fast Track Referral Model System be implemented. This network system helps to ensure that quality state-of-the-art technology is used, results are reported in a timely manner, and the tracking and fingerprinting of isolates of MTBC occur efficiently for optimal exchange of information between medical centers and the public health sector (54). CLSI guidelines recommend reporting results within 24 hours for acid-fast bacterium smears and interim reports if growth is seen on inoculated solid medium or when a positive signal is detected on liquid medium. A confirmatory report should be issued once growth is evident, and an identification of either NTM or MTBC should be reported. Moreover, reporting should be done as soon as species level identification is available for NTM and, finally, when susceptibility testing is completed (usually within 7 to 14 days for NTM with broth systems). Additionally, for MTBC, identification from culture results should be reported within an average of 14 days, and a complete culture report with susceptibility results should be reported within an average of 4 to 6 weeks. The use of liquid medium rapid detection

systems along with solid medium is critical in order to meet these guidelines (24, 54, 182). In an effort to develop a more consistent and practical method of reporting identifications performed by molecular techniques, the CLSI (54) has published interpretive criteria for the identification of mycobacterial species by DNA target sequencing, including consensus cutoff values for percent identity. The document states that mycobacteria with 100% 16S rRNA partial (500-bp) gene sequence identity for genus and species may be definitively reported with both genus and species. However, for those isolates whose identity is 99.0% to 99.9%, the recommendation is to report as “Mycobacterium most closely related to [the species given].” However, if the identity to a recognized species is ≥95 to 98.9%, the results should state that the isolate cannot be definitely identified by 16S rRNA gene sequencing; identification should be given as “most closely related to Mycobacterium sp.” Microheterogeneity within an NTM species (i.e., 1 to 5 bases in the 16S rRNA gene are different from the sequence of a known reference strain) has been described for several slowly growing species, including M. gordonae, M. bohemicum, M. kansasii, M. celatum, and M. lentiflavum. In general, members of the genus Mycobacterium are closely related to each other in their 16S rRNA gene sequences and may differ by only a few bases or not at all. In the absence of strong phenotypic differences, definitive NTM identification may be difficult, if not impossible. Turenne et al. advocate that the individual strain ambiguities should be examined carefully using optimum quality sequence databases for comparison, and even then, results may remain inconclusive without resorting to sequencing of alternate targets (89). As has been discussed, 16S rRNA partial gene sequencing is not useful for separation of some species, including the species within the MTBC. Thus, with MTBC, reporting results of only the 16S rRNA gene sequence should state “Mycobacterium tuberculosis complex.” When species level identification is necessary, alternate targets, such as gyrB, may provide species differentiation, except for MTB and M. canettii. Among the slowly growing NTM, several species are not identifiable with 16S rRNA gene sequencing alone. For example, M. kansasii and M. gastri share sequence identity in their complete 16S rRNA genes, and thus alternate gene targets or determination of photochromogenicity and/or growth rates are required to provide species resolution. Alternate gene targets that may help in cases where sequencing of the 16S rRNA gene is not adequate include the ITS 1 region, hsp65, gyrB, rpoB, recA, and dnaA genes. Specific numerical cutoff values such as are recommended for the 16S rRNA gene are not yet recommended for these genes. The more accurate characterization of new species by molecular technology and the enhancement of clinical data with standardized AST results continue to advance our knowledge of the MTBC, NTM, and disease caused by these species. Proper interpretation and reporting of species identification, even by molecular methods, should be checked against fundamentally established phenotypic characteristics, such as growth rate, colony morphology, and pigmentation. Determination of temperature requirements also may be useful for select isolates. Finally, appropriate AST of clinically significant species is vital to recognize and effectively treat multiple-drug-resistant mycobacterial species, including multidrug-resistant and extensively drug-resistant MTBC strains, as well as NTM such as M. simiae, MAC, rifampin-resistant M. kansasii, and other clinically significant drug-resistant species.

588 n

BACTERIOLOGY

REFERENCES 1. Tortoli E. 2003. Impact of genotypic studies on mycobacterial taxonomy: the new mycobacteria of the 1990s. Clin Microbiol Rev 16:319–354. 2. Ben Salah I, Cayrou C, Raoult D, Drancourt M. 2009. Mycobacterium marseillense sp. nov., Mycobacterium timonense sp. nov. and Mycobacterium bouchedurhonense sp. nov., members of the Mycobacterium avium complex. Int J Syst Evol Microbiol 59:2803–2808. 3. Hannigan GD, Krivogorsky B, Fordice D, Welch JB, Dahl JL. 2013. Mycobacterium minnesotense sp. nov., a photochromogenic bacterium isolated from sphagnum peat bogs. Int J Syst Evol Microbiol 63:124–128. 4. Kim BJ, Hong SH, Yu HK, Park YG, Jeong J, Lee SH, Kim SR, Kim K, Kook YH. 2012. Mycobacterium parakoreense sp. nov., a slowly growing non-chromogenic species related to Mycobacterium koreense, isolated from human clinical specimen. Int J Syst Evol Microbiol 63:2301–2308. 5. Kim BJ, Jeong J, Lee SH, Kim SR, Yu HK, Park YG, Kim KJ, Kook YH. 2012. Mycobacterium koreense sp. nov., a slowly growing non-chromogenic species closely related to Mycobacterium triviale. Int J Syst Evol Microbiol 62:1289– 1295. 6. Kim BJ, Math RK, Jeon CO, Yu HK, Park YG, Kook YH. 2012. Mycobacterium yongonense sp. nov., a novel slow growing nonchromogenic species closely related to Mycobacterium intracellulare. Int J Syst Evol Microbiol 63:192–199. 7. Lee HK, Lee SA, Lee IK, Yu HK, Park YG, Hyun JW, Kim K, Kook YH, Kim BJ. 2010. Mycobacterium paraseoulense sp. nov., a slowly growing, scotochromogenic species related genetically to Mycobacterium seoulense. Int J Syst Evol Microbiol 60:439–443. 8. Masaki T, Ohkusu K, Hata H, Fujiwara N, Iihara H, Yamada-Noda M, Nhung PH, Hayashi M, Asano Y, Kawamura Y, Ezaki T. 2006. Mycobacterium kumamotonense Sp. Nov. recovered from clinical specimen and the first isolation report of Mycobacterium arupense in Japan: novel slowly growing, nonchromogenic clinical isolates related to Mycobacterium terrae complex. Microbiol Immunol 50:889– 897. 9. Mun HS, Kim HJ, Oh EJ, Kim H, Bai GH, Yu HK, Park YG, Cha CY, Kook YH, Kim BJ. 2007. Mycobacterium seoulense sp. nov., a slowly growing scotochromogenic species. Int J Syst Evol Microbiol 57:594–599. 10. Mun HS, Park JH, Kim H, Yu HK, Park YG, Cha CY, Kook YH, Kim BJ. 2008. Mycobacterium senuense sp. nov., a slowly growing, non-chromogenic species closely related to the Mycobacterium terrae complex. Int J Syst Evol Microbiol 58:641–646. 11. Nakanaga K, Hoshino Y, Wakabayashi M, Fujimoto N, Tortoli E, Makino M, Tanaka T, Ishii N. 2012. Mycobacterium shigaense sp. nov., a novel slowly growing scotochromogenic mycobacterium that produced nodules in an erythroderma patient with severe cellular immunodeficiency and a history of Hodgkin’s disease. J Dermatol 39:389–396. 12. Okazaki M, Ohkusu K, Hata H, Ohnishi H, Sugahara K, Kawamura C, Fujiwara N, Matsumoto S, Nishiuchi Y, Toyoda K, Saito H, Yonetani S, Fukugawa Y, Yamamoto M, Wada H, Sejimo A, Ebina A, Goto H, Ezaki T, Watanabe T. 2009. Mycobacterium kyorinense sp. nov., a novel, slow-growing species, related to Mycobacterium celatum, isolated from human clinical specimens. Int J Syst Evol Microbiol 59:1336–1341. 13. Pourahmad F, Cervellione F, Thompson KD, Taggart JB, Adams A, Richards RH. 2008. Mycobacterium stomatepiae sp. nov., a slowly growing, non-chromogenic species isolated from fish. Int J Syst Evol Microbiol 58:2821–2827.

14. Saito H, Iwamoto T, Ohkusu K, Otsuka Y, Akiyama Y, Sato S, Taguchi O, Sueyasu Y, Kawabe Y, Fujimoto H, Ezaki T, Butler R. 2011. Mycobacterium shinjukuense sp. nov., a slowly growing, non-chromogenic species isolated from human clinical specimens. Int J Syst Evol Microbiol 61:1927–1932. 15. Tortoli E, Bottger EC, Fabio A, Falsen E, Gitti Z, Grottola A, Klenk HP, Mannino R, Mariottini A, Messino M, Pecorari M, Rumpianesi F. 2011. Mycobacterium europaeum sp. nov., a scotochromogenic species related to the Mycobacterium simiae complex. Int J Syst Evol Microbiol 61:1606–1611. 16. Tortoli E, Gitti Z, Klenk HP, Lauria S, Mannino R, Mantegani P, Mariottini A, Neonakis I. 2013. Survey of 150 strains belonging to the Mycobacterium terrae complex and description of Mycobacterium engbaekii sp. nov., Mycobacterium heraklionense sp. nov. and Mycobacterium longobardum sp. nov. Int J Syst Evol Microbiol 63:401–411. 17. van Ingen J, Al-Hajoj SA, Boeree M, Al-Rabiah F, Enaimi M, de Zwaan R, Tortoli E, Dekhuijzen R, van Soolingen D. 2009. Mycobacterium riyadhense sp. nov., a non-tuberculous species identified as Mycobacterium tuberculosis complex by a commercial line-probe assay. Int J Syst Evol Microbiol 59:1049–1053. 18. van Ingen J, Boeree MJ, de Lange WC, de Haas PE, van der Zanden AG, Mijs W, Rigouts L, Dekhuijzen PN, van Soolingen D. 2009. Mycobacterium noviomagense sp. nov.; clinical relevance evaluated in 17 patients. Int J Syst Evol Microbiol 59:845–849. 19. van Ingen J, Lindeboom JA, Hartwig NG, de Zwaan R, Tortoli E, Dekhuijzen PN, Boeree MJ, van Soolingen D. 2009. Mycobacterium mantenii sp. nov., a pathogenic, slowly growing, scotochromogenic species. Int J Syst Evol Microbiol 59:2782–2787. 20. van Ingen J, Tortoli E, Selvarangan R, Coyle MB, Crump JA, Morrissey AB, Dekhuijzen PN, Boeree MJ, van Soolingen D. 2011. Mycobacterium sherrisii sp. nov., a slowgrowing non-chromogenic species. Int J Syst Evol Microbiol 61:1293–1298. 21. Kim BJ, Hong SH, Kook YH. 2014. Mycobacterium paragordonae sp. nov., a slowly growing, scotochromogenic species closely related to Mycobacterium gordonae. Int J Syst Evol Microbiol 64:39–45. 22. Ramos JP, Campos CE, Caldas PC, Ferreira NV, da Silva MV, Redner P, Campelo CL, Vale SF, Barroso EC, Medeiros RF, Montes FC, Galvao TC, Tortoli E. 2013. Mycobacterium fragae sp. nov., a non-chromogenic species isolated from human respiratory specimens. Int J Syst Evol Microbiol 63:2583–2587. 23. Sahraoui N, Ballif M, Zelleg S, Yousfi N, Ritter C, Friedel U, Amstutz B, Yala D, Boulahbal F, Guetarni D, Zinsstag J, Keller PM. 2011. Mycobacterium algericum sp. nov., a novel rapidly growing species related to the Mycobacterium terrae complex and associated with goat lung lesions. Int J Syst Evol Microbiol 61:1870–1874. 24. Griffith DE, Aksamit T, Brown-Elliott BA, Catanzaro A, Daley C, Gordin F, Holland SM, Horsburgh R, Huitt G, Iademarco MF, Iseman M, Olivier K, Ruoss S, von Reyn CF, Wallace RJ Jr, Winthrop K. 2007. An official ATS/IDSA statement: diagnosis, treatment, and prevention of nontuberculous mycobacterial diseases. Am J Respir Crit Care Med 175:367–416. 25. Marras TK, Chedore P, Ying AM, Jamieson F. 2007. Isolation prevalence of pulmonary non-tuberculous mycobacteria in Ontario, 1997–2003. Thorax 62:661–666. 26. Martin-Casabona N, Bahrmand AR, Bennedsen J, Thomsen VO, Curcio M, Fauville-Dufaux M, Feldman K, Havelkova M, Katila ML, Koksalan K, Pereira MF, Rodrigues F, Pfyffer GE, Portaels F, Urgell JR, RuschGerdes S, Tortoli E, Vincent V, Watt B. 2004. Non-

31. Characteristics of Slowly Growing Mycobacteria n 589

27.

28.

29.

30.

31.

32.

33.

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

tuberculous mycobacteria: patterns of isolation. A multicountry retrospective survey. Int J Tuberc Lung Dis 8:1186– 1193. Good RC. 1980. From the Center for Disease Control. Isolation of nontuberculous mycobacteria in the United States, 1979. J Infect Dis 142:779–783. O’Brien RJ, Geiter LJ, Snider DE Jr. 1987. The epidemiology of nontuberculous mycobacterial diseases in the United States. Results from a national survey. Am Rev Respir Dis 135:1007–1014. Horsburgh CR Jr, Selik RM. 1989. The epidemiology of disseminated nontuberculous mycobacterial infection in the acquired immunodeficiency syndrome (AIDS). Am Rev Respir Dis 139:4–7. Cassidy PM, Hedberg K, Saulson A, McNelly E, Winthrop KL. 2009. Nontuberculous mycobacterial disease prevalence and risk factors: a changing epidemiology. Clin Infect Dis 49:e124–e129. Behr MA, Falkinham JO III. 2009. Molecular epidemiology of nontuberculous mycobacteria. Future Microbiol 4:1009–1020. Lawn SD, Edwards DJ, Kranzer K, Vogt M, Bekker LG, Wood R. 2009. Urine lipoarabinomannan assay for tuberculosis screening before antiretroviral therapy diagnostic yield and association with immune reconstitution disease. AIDS 23:1875–1880. Minion J, Leung E, Talbot E, Dheda K, Pai M, Menzies D. 2011. Diagnosing tuberculosis with urine lipoarabinomannan: systematic review and meta-analysis. Eur Respir J 38:1398–1405. Adékambi T, Colson P, Drancourt M. 2003. rpoB-based identification of nonpigmented and late-pigmenting rapidly growing mycobacteria. J Clin Microbiol 41:5699–5708. Daley P, Thomas S, Pai M. 2007. Nucleic acid amplification tests for the diagnosis of tuberculous lymphadenitis: a systematic review. Int J Tuberc Lung Dis 11:1166–1176. Dinnes J, Deeks J, Kunst H, Gibson A, Cummins E, Waugh N, Drobniewski F, Lalvani A. 2007. A systematic review of rapid diagnostic tests for the detection of tuberculosis infection. Health Technol Assess 11:1–196. Flores LL, Pai M, Colford JM Jr, Riley LW. 2005. Inhouse nucleic acid amplification tests for the detection of Mycobacterium tuberculosis in sputum specimens: metaanalysis and meta-regression. BMC Microbiol 5:55. Ling DI, Flores LL, Riley LW, Pai M. 2008. Commercial nucleic-acid amplification tests for diagnosis of pulmonary tuberculosis in respiratory specimens: meta-analysis and meta-regression. PLoS One 3:e1536. Pai M, Flores LL, Hubbard A, Riley LW, Colford JM Jr. 2004. Nucleic acid amplification tests in the diagnosis of tuberculous pleuritis: a systematic review and metaanalysis. BMC Infect Dis 4:6. Pai M, Kalantri S, Dheda K. 2006. New tools and emerging technologies for the diagnosis of tuberculosis: part II. Active tuberculosis and drug resistance. Expert Rev Mol Diagn 6:423–432. Park H, Jang H, Kim C, Chung B, Chang CL, Park SK, Song S. 2000. Detection and identification of mycobacteria by amplification of the internal transcribed spacer regions with genus- and species-specific PCR primers. J Clin Microbiol 38:4080–4085. Piersimoni C, Scarparo C. 2003. Relevance of commercial amplification methods for direct detection of Mycobacterium tuberculosis complex in clinical samples. J Clin Microbiol 41:5355–5365. CDC. 2009. Updated guidelines for the use of nucleic acid amplification tests in the diagnosis of tuberculosis. MMWR Morb Mortal Wkly Rep 58:7–10. CDC. 2013. Availability of an assay for detecting Mycobacterium tuberculosis, including rifampin-resistant strains, and

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

considerations for its use—United States, 2013. MMWR Morb Mortal Wkly Rep 62:821–827. Mehta PK, Raj A, Singh N, Khuller GK. 2012. Diagnosis of extrapulmonary tuberculosis by PCR. FEMS Immunol Med Microbiol 66:20–36. Pai M, Flores LL, Pai N, Hubbard A, Riley LW, Colford JM Jr. 2003. Diagnostic accuracy of nucleic acid amplification tests for tuberculous meningitis: a systematic review and meta-analysis. Lancet Infect Dis 3:633–643. Yang YC, Lu PL, Huang SC, Jenh YS, Jou R, Chang TC. 2011. Evaluation of the Cobas TaqMan MTB test for direct detection of Mycobacterium tuberculosis complex in respiratory specimens. J Clin Microbiol 49:797–801. Lawn SD, Nicol MP. 2011. Xpert(R) MTB/RIF assay: development, evaluation and implementation of a new rapid molecular diagnostic for tuberculosis and rifampicin resistance. Future Microbiol 6:1067–1082. Tortoli E, Russo C, Piersimoni C, Mazzola E, Dal Monte P, Pascarella M, Borroni E, Mondo A, Piana F, Scarparo C, Coltella L, Lombardi G, Cirillo DM. 2012. Clinical validation of Xpert MTB/RIF for the diagnosis of extrapulmonary tuberculosis. Eur Respir J 40:442–447. Vadwai V, Boehme C, Nabeta P, Shetty A, Alland D, Rodrigues C. 2011. Xpert MTB/RIF: a new pillar in diagnosis of extrapulmonary tuberculosis? J Clin Microbiol 49:2540–2545. WHO. 2011, posting date. Rapid implementation of the Xpert MTB/RIF diagnostic test, 2011. http://whqlibdoc. who.int/publications/2011/9789241501569_eng.pdf. Van Rie A, Mellet K, John MA, Scott L, Page-Shipp L, Dansey H, Victor T, Warren R. 2012. False-positive rifampicin resistance on Xpert(R) MTB/RIF: case report and clinical implications. Int J Tuberc Lung Dis 16:206– 208. Williamson DA, Basu I, Bower J, Freeman JT, Henderson G, Roberts SA. 2012. An evaluation of the Xpert MTB/RIF assay and detection of false-positive rifampicin resistance in Mycobacterium tuberculosis. Diagn Microbiol Infect Dis 74:207–209. Clinical and Laboratory Standards Institute. 2008. Interpretive Criteria for Identification of Bacteria and Fungi by DNA Target Sequencing; Approved Guideline. MM18-A. CLSI, Wayne, PA. Clinical and Laboratory Standards Institute. 2008. Laboratory Detection and Identification of Mycobacteria; Approved Guideline. M48-A. CLSI, Wayne, PA. Alshamaony L, Goodfellow M, Minnikin DE, Mordarska H. 1976. Free mycolic acids as criteria in the classification of Gordona and the “rhodochrous” complex. J Gen Microbiol 92:183–187. Butler WR, Guthertz LS. 2001. Mycolic acid analysis by high-performance liquid chromatography for identification of Mycobacterium species. Clin Microbiol Rev 14:704–726. Lechevalier MP, Horan AC, Lechevalier H. 1971. Lipid composition in the classification of nocardiae and mycobacteria. J Bacteriol 105:313–318. Fox GE, Stackebrandt E, Hespell RB, Gibson J, Maniloff J, Dyer TA, Wolfe RS, Balch WE, Tanner RS, Magrum LJ, Zablen LB, Blakemore R, Gupta R, Bonen L, Lewis BJ, Stahl DA, Luehrsen KR, Chen KN, Woese CR. 1980. The phylogeny of prokaryotes. Science 209:457–463. Kim BJ, Lee SH, Lyu MA, Kim SJ, Bai GH, Chae GT, Kim EC, Cha CY, Kook YH. 1999. Identification of mycobacterial species by comparative sequence analysis of the RNA polymerase gene (rpoB). J Clin Microbiol 37:1714– 1720. Kirschner P, Springer B, Vogel U, Meier A, Wrede A, Kiekenbeck M, Bange FC, Bottger EC. 1993. Genotypic identification of mycobacteria by nucleic acid sequence determination: report of a 2-year experience in a clinical laboratory. J Clin Microbiol 31:2882–2889.

590 n

BACTERIOLOGY

62. Lane DJ, Pace B, Olsen GJ, Stahl DA, Sogin ML, Pace NR. 1985. Rapid determination of 16S ribosomal RNA sequences for phylogenetic analyses. Proc Natl Acad Sci USA 82:6955–6959. 63. McNabb A, Eisler D, Adie K, Amos M, Rodrigues M, Stephens G, Black WA, Isaac-Renton J. 2004. Assessment of partial sequencing of the 65-kilodalton heat shock protein gene (hsp65) for routine identification of Mycobacterium species isolated from clinical sources. J Clin Microbiol 42:3000–3011. 64. Plikaytis BB, Plikaytis BD, Yakrus MA, Butler WR, Woodley CL, Silcox VA, Shinnick TM. 1992. Differentiation of slowly growing Mycobacterium species, including Mycobacterium tuberculosis, by gene amplification and restriction fragment length polymorphism analysis. J Clin Microbiol 30:1815–1822. 65. Roth A, Fischer M, Hamid ME, Michalke S, Ludwig W, Mauch H. 1998. Differentiation of phylogenetically related slowly growing mycobacteria based on 16S-23S rRNA Gene internal transcribed spacer sequences. J Clin Microbiol 36:139–147. 66. Williams KJ, Ling CL, Jenkins C, Gillespie SH, McHugh TD. 2007. A paradigm for the molecular identification of Mycobacterium species in a routine diagnostic laboratory. J Med Microbiol 56:598–602. 67. Boddinghaus B, Rogall T, Flohr T, Blocker H, Bottger EC. 1990. Detection and identification of mycobacteria by amplification of rRNA. J Clin Microbiol 28:1751–1759. 68. Cook VJ, Turenne CY, Wolfe J, Pauls R, Kabani A. 2003. Conventional methods versus 16S ribosomal DNA sequencing for identification of nontuberculous mycobacteria: cost analysis. J Clin Microbiol 41:1010–1015. 69. El Amin NM, Hanson HS, Pettersson B, Petrini B, Von Stedingk LV. 2000. Identification of non-tuberculous mycobacteria: 16S rRNA gene sequence analysis vs. conventional methods. Scand J Infect Dis 32:47–50. 70. Han XY, Pham AS, Tarrand JJ, Sood PK, Luthra R. 2002. Rapid and accurate identification of mycobacteria by sequencing hypervariable regions of the 16S ribosomal RNA gene. Am J Clin Pathol 118:796–801. 71. Holberg-Petersen M, Steinbakk M, Figenschau KJ, Jantzen E, Eng J, Melby KK. 1999. Identification of clinical isolates of Mycobacterium spp. by sequence analysis of the 16S ribosomal RNA gene. Experience from a clinical laboratory. APMIS 107:231–239. 72. Pauls RJ, Turenne CY, Wolfe JN, Kabani A. 2003. A high proportion of novel mycobacteria species identified by 16S rDNA analysis among slowly growing AccuProbenegative strains in a clinical setting. Am J Clin Pathol 120:560–566. 73. Springer B, Stockman L, Teschner K, Roberts GD, Bottger EC. 1996. Two-laboratory collaborative study on identification of mycobacteria: molecular versus phenotypic methods. J Clin Microbiol 34:296–303. 74. Woo PCY, Lau SKP, Teng JLL, Tse H, Yuen KY. 2008. Then and now: use of 16S rDNA gene sequencing for bacterial identification and discovery of novel bacteria in clinical microbiology laboratories. Clin Microbiol Infect 14:908–934. 75. Kirschner P, Bottger EC. 1992. Microheterogeneity within rRNA of Mycobacterium gordonae. J Clin Microbiol 30:1049–1050. 76. Richter E, Niemann S, Rusch-Gerdes S, Hoffner S. 1999. Identification of Mycobacterium kansasii by using a DNA probe (AccuProbe) and molecular techniques. J Clin Microbiol 37:964–970. 77. Springer B, Wu WK, Bodmer T, Haase G, Pfyffer GE, Kroppenstedt RM, Schroder KH, Emler S, Kilburn JO, Kirschner P, Telenti A, Coyle MB, Bottger EC. 1996. Isolation and characterization of a unique group of slowly

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

89.

90.

91.

92.

93.

growing mycobacteria: description of Mycobacterium lentiflavum sp. nov. J Clin Microbiol 34:1100–1107. Liesack W, Sela S, Bercovier H, Pitulle C, Stackebrandt E. 1991. Complete nucleotide sequence of the Mycobacterium leprae 23 S and 5 S rRNA genes plus flanking regions and their potential in designing diagnostic oligonucleotide probes. FEBS Lett 281:114–118. van der Giessen JW, Haring RM, van der Zeijst BA. 1994. Comparison of the 23S ribosomal RNA genes and the spacer region between the 16S and 23S rRNA genes of the closely related Mycobacterium avium and Mycobacterium paratuberculosis and the fast-growing Mycobacterium phlei. Microbiology 140(Part 5):1103–1108. Harmsen D, Dostal S, Roth A, Niemann S, Rothganger J, Sammeth M, Albert J, Frosch M, Richter E. 2003. RIDOM: comprehensive and public sequence database for identification of Mycobacterium species. BMC Infect Dis 3:26. Roth A, Reischl U, Streubel A, Naumann L, Kroppenstedt RM, Habicht M, Fischer M, Mauch H. 2000. Novel diagnostic algorithm for identification of mycobacteria using genus-specific amplification of the 16S-23S rRNA gene spacer and restriction endonucleases. J Clin Microbiol 38:1094–1104. Frothingham R, Wilson KH. 1993. Sequence-based differentiation of strains in the Mycobacterium avium complex. J Bacteriol 175:2818–2825. Frothingham R, Wilson KH. 1994. Molecular phylogeny of the Mycobacterium avium complex demonstrates clinically meaningful divisions. J Infect Dis 169:305–312. Mohamed AM, Kuyper DJ, Iwen PC, Ali HH, Bastola DR, Hinrichs SH. 2005. Computational approach involving use of the internal transcribed spacer 1 region for identification of Mycobacterium species. J Clin Microbiol 43:3811– 3817. Devallois A, Goh KS, Rastogi N. 1997. Rapid identification of mycobacteria to species level by PCR-restriction fragment length polymorphism analysis of the hsp65 gene and proposition of an algorithm to differentiate 34 mycobacterial species. J Clin Microbiol 35:2969–2973. Telenti A, Marchesi F, Balz M, Bally F, Bottger EC, Bodmer T. 1993. Rapid identification of mycobacteria to the species level by polymerase chain reaction and restriction enzyme analysis. J Clin Microbiol 31:175–178. Ringuet H, Akoua-Koffi C, Honore S, Varnerot A, Vincent V, Berche P, Gaillard JL, Pierre-Audigier C. 1999. hsp65 Sequencing for identification of rapidly growing mycobacteria. J Clin Microbiol 37:852–857. Lee H, Bang HE, Bai GH, Cho SN. 2003. Novel polymorphic region of the rpoB gene containing Mycobacterium species-specific sequences and its use in identification of mycobacteria. J Clin Microbiol 41:2213–2218. Turenne CY, Tschetter L, Wolfe J, Kabani A. 2001. Necessity of quality-controlled 16S rRNA gene sequence databases: identifying nontuberculous Mycobacterium species. J Clin Microbiol 39:3637–3648. Tuohy MJ, Hall GS, Sholtis M, Procop GW. 2005. Pyrosequencing as a tool for the identification of common isolates of Mycobacterium sp. Diagn Microbiol Infect Dis 51:245–250. Heller LC, Jones M, Widen RH. 2008. Comparison of DNA pyrosequencing with alternative methods for identification of mycobacteria. J Clin Microbiol 46:2092–2094. Bao JR, Master RN, Schwab DA, Clark RB. 2010. Identification of acid-fast bacilli using pyrosequencing analysis. Diagn Microbiol Infect Dis 67:234–238. Wengenack NL, Hall L. 2011. Sequence-based identification and characterization of mycobacteria, p 437–452. In Persing DH (ed), Molecular Microbiology: Diagnostic Principles and Practice, 2nd ed. ASM Press, Washington, DC.

31. Characteristics of Slowly Growing Mycobacteria n 591 94. Kasai H, Ezaki T, Harayama S. 2000. Differentiation of phylogenetically related slowly growing mycobacteria by their gyrB sequences. J Clin Microbiol 38:301–308. 95. Niemann S, Harmsen D, Rusch-Gerdes S, Richter E. 2000. Differentiation of clinical Mycobacterium tuberculosis complex isolates by gyrB DNA sequence polymorphism analysis. J Clin Microbiol 38:3231–3234. 96. Pinsky BA, Banaei N. 2008. Multiplex real-time PCR assay for rapid identification of Mycobacterium tuberculosis complex members to the species level. J Clin Microbiol 46:2241–2246. 97. Somoskovi A, Dormandy J, Parsons LM, Kaswa M, Goh KS, Rastogi N, Salfinger M. 2007. Sequencing of the pncA gene in members of the Mycobacterium tuberculosis complex has important diagnostic applications: identification of a species-specific pncA mutation in “Mycobacterium canettii” and the reliable and rapid predictor of pyrazinamide resistance. J Clin Microbiol 45:595–599. 98. Neonakis IK, Gitti Z, Petinaki E, Maraki S, Spandidos DA. 2007. Evaluation of the GenoType MTBC assay for differentiating 120 clinical Mycobacterium tuberculosis complex isolates. Eur J Clin Microbiol Infect Dis 26:151– 152. 99. Richter E, Weizenegger M, Fahr AM, Rusch-Gerdes S. 2004. Usefulness of the GenoType MTBC assay for differentiating species of the Mycobacterium tuberculosis complex in cultures obtained from clinical specimens. J Clin Microbiol 42:4303–4306. 100. Richter E, Weizenegger M, Rusch-Gerdes S, Niemann S. 2003. Evaluation of genotype MTBC assay for differentiation of clinical Mycobacterium tuberculosis complex isolates. J Clin Microbiol 41:2672–2675. 101. Somoskovi A, Dormandy J, Rivenburg J, Pedrosa M, McBride M, Salfinger M. 2008. Direct comparison of the genotype MTBC and genomic deletion assays in terms of ability to distinguish between members of the Mycobacterium tuberculosis complex in clinical isolates and in clinical specimens. J Clin Microbiol 46:1854–1857. 102. Halse TA, Escuyer VE, Musser KA. 2011. Evaluation of a single-tube multiplex real-time PCR for differentiation of members of the Mycobacterium tuberculosis complex in clinical specimens. J Clin Microbiol 49:2562–2567. 103. Bull TJ, Shanson DC. 1992. Evaluation of a commercial chemiluminescent gene probe system ‘AccuProbe’ for the rapid differentiation of mycobacteria, including ‘MAIC X’, isolated from blood and other sites, from patients with AIDS. J Hosp Infect 21:143–149. 104. Lebrun L, Espinasse F, Poveda JD, Vincent-Levy-Frebault V. 1992. Evaluation of nonradioactive DNA probes for identification of mycobacteria. J Clin Microbiol 30:2476–2478. 105. Middleton AM, Chadwick MV, Gaya H. 1997. Detection of Mycobacterium tuberculosis in mixed broth cultures using DNA probes. Clin Microbiol Infect 3:668–671. 106. Tenover FC, Crawford JT, Huebner RE, Geiter LJ, Horsburgh CR Jr, Good RC. 1993. The resurgence of tuberculosis: is your laboratory ready? J Clin Microbiol 31:767–770. 107. Tortoli E, Simonetti MT, Lavinia F. 1996. Evaluation of reformulated chemiluminescent DNA probe (AccuProbe) for culture identification of Mycobacterium kansasii. J Clin Microbiol 34:2838–2840. 108. Cloud JL, Carroll KC, Cohen S, Anderson CM, Woods GL. 2005. Interpretive criteria for use of AccuProbe for identification of Mycobacterium avium complex directly from 7H9 broth cultures. J Clin Microbiol 43:3474–3478. 109. Tortoli E, Pecorari M, Fabio G, Messino M, Fabio A. 2010. Commercial DNA probes for mycobacteria incorrectly identify a number of less frequently encountered species. J Clin Microbiol 48:307–310.

110. Christiansen DC, Roberts GD, Patel R. 2004. Mycobacterium celatum, an emerging pathogen and cause of false positive amplified Mycobacterium tuberculosis direct test. Diagn Microbiol Infect Dis 49:19–24. 111. Richter E, Niemann S, Gloeckner FO, Pfyffer GE, Rusch-Gerdes S. 2002. Mycobacterium holsaticum sp. nov. Int J Syst Evol Microbiol 52:1991–1996. 112. Daley P, Petrich A, May K, Luinstra K, Rutherford C, Chedore P, Jamieson F, Smieja M. 2008. Comparison of in-house and commercial 16S rRNA sequencing with high-performance liquid chromatography and genotype AS and CM for identification of nontuberculous mycobacteria. Diagn Microbiol Infect Dis 61:284–293. 113. Gitti Z, Neonakis I, Fanti G, Kontos F, Maraki S, Tselentis Y. 2006. Use of the GenoType Mycobacterium CM and AS assays to analyze 76 nontuberculous mycobacterial isolates from Greece. J Clin Microbiol 44:2244–2246. 114. Lee AS, Jelfs P, Sintchenko V, Gilbert GL. 2009. Identification of non-tuberculous mycobacteria: utility of the GenoType Mycobacterium CM/AS assay compared with HPLC and 16S rRNA gene sequencing. J Med Microbiol 58:900–904. 115. Makinen J, Sarkola A, Marjamaki M, Viljanen MK, Soini H. 2002. Evaluation of genotype and LiPA MYCOBACTERIA assays for identification of Finnish mycobacterial isolates. J Clin Microbiol 40:3478–3481. 116. Richter E, Rusch-Gerdes S, Hillemann D. 2006. Evaluation of the GenoType Mycobacterium Assay for identification of mycobacterial species from cultures. J Clin Microbiol 44:1769–1775. 117. Russo C, Tortoli E, Menichella D. 2006. Evaluation of the new GenoType Mycobacterium assay for identification of mycobacterial species. J Clin Microbiol 44:334– 339. 118. Tortoli E, Mariottini A, Mazzarelli G. 2003. Evaluation of INNO-LiPA MYCOBACTERIA v2: improved reverse hybridization multiple DNA probe assay for mycobacterial identification. J Clin Microbiol 41:4418–4420. 119. Hofmann-Thiel S, Turaev L, Alnour T, Drath L, Mullerova M, Hoffmann H. 2011. Multi-centre evaluation of the speed-oligo Mycobacteria assay for differentiation of Mycobacterium spp. in clinical isolates. BMC Infect Dis 11:353. 120. Quezel-Guerraz NM, Arriaza MM, Avila JA, SanchezYebra Romera WE, Martinez-Lirola MJ. 2010. Evaluation of the Speed-oligo(R) Mycobacteria assay for identification of Mycobacterium spp. from fresh liquid and solid cultures of human clinical samples. Diagn Microbiol Infect Dis 68:123–131. 121. Lara-Oya A, Mendoza-Lopez P, Rodriguez-Granger J, Fernandez-Sanchez AM, Bermudez-Ruiz MP, Toro-Peinado I, Palop-Borras B, Navarro-Mari JM, MartinezLirola MJ. 2013. Evaluation of the speed-oligo direct Mycobacterium tuberculosis assay for molecular detection of mycobacteria in clinical respiratory specimens. J Clin Microbiol 51:77–82. 122. Richardson ET, Samson D, Banaei N. 2009. Rapid identification of Mycobacterium tuberculosis and nontuberculous mycobacteria by multiplex, real-time PCR. J Clin Microbiol 47:1497–1502. 123. Halse TA, Escuyer VE, Musser KA. 2011. Evaluation of a single-tube multiplex real-time PCR for differentiation of members of the Mycobacterium tuberculosis complex in clinical specimens. J Clin Microbiol 49:2562–2567. 124. Pinsky BA, Banaei N. 2008. Multiplex real-time PCR assay for rapid identification of Mycobacterium tuberculosis complex members to the species level. J Clin Microbiol 46:2241–2246. 125. Liu Q, Luo T, Li J, Mei J, Gao Q. 2013. Triplex real-time PCR melting curve analysis for detecting Mycobacterium

592 n

126.

127.

128.

129.

130.

131.

132.

133.

134.

135.

136.

137.

138.

139.

BACTERIOLOGY

tuberculosis mutations associated with resistance to second-line drugs in a single reaction. J Antimicrob Chemother 68:1097–1103. Luo T, Jiang L, Sun W, Fu G, Mei J, Gao Q. 2011. Multiplex real-time PCR melting curve assay to detect drug-resistant mutations of Mycobacterium tuberculosis. J Clin Microbiol 49:3132–3138. Martinez AN, Britto CF, Nery JA, Sampaio EP, Jardim MR, Sarno EN, Moraes MO. 2006. Evaluation of realtime and conventional PCR targeting complex 85 genes for detection of Mycobacterium leprae DNA in skin biopsy samples from patients diagnosed with leprosy. J Clin Microbiol 44:3154–3159. Parkash O, Singh HB, Rai S, Pandey A, Katoch VM, Girdhar BK. 2004. Detection of Mycobacterium leprae DNA for 36kDa protein in urine from leprosy patients: a preliminary report. Rev Inst Med Trop Sao Paulo 46:275– 277. Patrocinio LG, Goulart IM, Goulart LR, Patrocinio JA, Ferreira FR, Fleury RN. 2005. Detection of Mycobacterium leprae in nasal mucosa biopsies by the polymerase chain reaction. FEMS Immunol Med Microbiol 44:311– 316. Phetsuksiri B, Rudeeaneksin J, Supapkul P, Wachapong S, Mahotarn K, Brennan PJ. 2006. A simplified reverse transcriptase PCR for rapid detection of Mycobacterium leprae in skin specimens. FEMS Immunol Med Microbiol 48:319–328. Santos AR, Balassiano V, Oliveira ML, Pereira MA, Santos PB, Degrave WM, Suffys PN. 2001. Detection of Mycobacterium leprae DNA by polymerase chain reaction in the blood of individuals, eight years after completion of anti-leprosy therapy. Mem Inst Oswaldo Cruz 96:1129–1133. Torres P, Camarena JJ, Gomez JR, Nogueira JM, Gimeno V, Navarro JC, Olmos A. 2003. Comparison of PCR mediated amplification of DNA and the classical methods for detection of Mycobacterium leprae in different types of clinical samples in leprosy patients and contacts. Lepr Rev 74:18–30. Chae GT, Kim MJ, Kang TJ, Lee SB, Shin HK, Kim JP, Ko YH, Kim SH, Kim NH. 2002. DNA-PCR and RT-PCR for the 18-kDa gene of Mycobacterium leprae to assess the efficacy of multi-drug therapy for leprosy. J Med Microbiol 51:417–422. Donoghue HD, Holton J, Spigelman M. 2001. PCR primers that can detect low levels of Mycobacterium leprae DNA. J Med Microbiol 50:177–182. Hartskeerl RA, de Wit MY, Klatser PR. 1989. Polymerase chain reaction for the detection of Mycobacterium leprae. J Gen Microbiol 135:2357–2364. Plikaytis BB, Gelber RH, Shinnick TM. 1990. Rapid and sensitive detection of Mycobacterium leprae using a nested-primer gene amplification assay. J Clin Microbiol 28:1913–1917. Bang PD, Suzuki K, Phuong le T, Chu TM, Ishii N, Khang TH. 2009. Evaluation of polymerase chain reaction-based detection of Mycobacterium leprae for the diagnosis of leprosy. J Dermatol 36:269–276. Lotz A, Ferroni A, Beretti J-L, Dauphin B, Carbonnelle E, Guet-Revillet H, Veziris N, Heym B, Jarlier V, Gaillard J-L, Pierre-Audigier C, Frapy E, Berche P, Nassif X, Bille E. 2010. Rapid identification of mycobacterial whole cells in solid and liquid culture media by matrixassisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 48:4481–4486. Saleeb PG, Drake SK, Murray PR, Zelazny AM. 2011. Identification of mycobacteria in solid-culture media by matrix-assisted laser desorption ionization–time of flight mass spectrometry. J Clin Microbiol 49:1790–1794.

140. El Khéchine A, Couderc C, Flaudrops C, Raoult D, Drancourt M. 2011. Matrix-assisted laser desorption/ionization time-of-flight mass spectrometry identification of mycobacteria in routine clinical practice. PLoS One 6:e24720. 141. Buckwalter SP, Connelly BJ, Olson SL, Lucas BC, Rodning AA, Doerr KA, Deml SM, Wohlfiel SL, Wengenack NL. 2013. Identification of Mycobacteria using 16S sequencing and MALDI-TOF MS, abstr C-3181. Abstr. 113th Gen. Meet. Am. Soc. Microbiol., Denver, CO. 142. Saleeb PG, Drake SK, Murray PR, Zelazny AM. 2011. Identification of mycobacteria in solid-culture media by matrix-assisted laser desorption ionization-time of flight mass spectrometry. J Clin Microbiol 49:1790–1794. 143. Mather CA, Rivera SF, Butler-Wu SM. 2014. Comparison of the Bruker Biotyper and Vitek MS matrix-assisted laser desorption ionization-time of flight mass spectrometry systems for identification of mycobacteria using simplified protein extraction protocols. J Clin Microbiol 52:130– 138. 144. Brown-Elliott BA, Wallace RJ. 2012. Enhancement of conventional phenotypic methods with molecular-based methods for the more definitive identification of nontuberculous mycobacteria. Clin Microbiol Newsl 34:109– 116. 145. Dhiman N, Hall L, Wohlfiel SL, Buckwalter SP, Wengenack NL. 2011. Performance and cost analysis of matrix-assisted laser desorption ionization-time of flight mass spectrometry for routine identification of yeast. J Clin Microbiol 49:1614–1616. 146. Massire C, Ivy CA, Lovari R, Kurepina N, Li H, Blyn LB, Hofstadler SA, Khechinashvili G, Stratton CW, Sampath R, Tang Y-W, Ecker DJ, Kreiswirth BN. 2011. Simultaneous identification of mycobacterial isolates to the species level and determination of tuberculosis drug resistance by PCR followed by electrospray ionization mass spectrometry. J Clin Microbiol 49:908–917. 147. Wang F, Massire C, Li H, Cummins LL, Li F, Jin J, Fan X, Wang S, Shao L, Zhang S, Meng S, Wu J, Lu C, Blyn LB, Sampath R, Ecker DJ, Zhang W, Tang Y-W. 2011. Molecular characterization of drug-resistant Mycobacterium tuberculosis isolates circulating in China by multilocus PCR and electrospray ionization mass spectrometry. J Clin Microbiol 49:2719–2721. 148. Simner PJ, Buckwalter S, Wengenack NL. 2013. Identification of Mycobacteria spp and Mycobacterium tuberculosis complex resistance determinants using the MDR-TB assay on the PLEX-ID PCR-electrospray ionization mass spectrometry (ESI-MS) System. Canadian Association of Clinical Microbiology and Infectious Diseases Annual Meeting, Quebec City, QC, Canada. 149. Yakrus MA, Reeves MW, Hunter SB. 1992. Characterization of isolates of Mycobacterium avium serotypes 4 and 8 from patients with AIDS by multilocus enzyme electrophoresis. J Clin Microbiol 30:1474–1478. 150. Cave MD, Murray M, Nardell E. 2005. Molecular epidemiology of Mycobacterium tuberculosis. ASM Press, Washington, DC. 151. van Embden JD, Cave MD, Crawford JT, Dale JW, Eisenach KD, Gicquel B, Hermans P, Martin C, McAdam R, Shinnick TM. 1993. Strain identification of Mycobacterium tuberculosis by DNA fingerprinting: recommendations for a standardized methodology. J Clin Microbiol 31:406–409. 152. Kamerbeek J, Schouls L, Kolk A, van Agterveld M, van Soolingen D, Kuijper S, Bunschoten A, Molhuizen H, Shaw R, Goyal M, van Embden J. 1997. Simultaneous detection and strain differentiation of Mycobacterium tuberculosis for diagnosis and epidemiology. J Clin Microbiol 35:907–914.

31. Characteristics of Slowly Growing Mycobacteria n 593 153. Supply P, Allix C, Lesjean S, Cardoso-Oelemann M, Rusch-Gerdes S, Willery E, Savine E, de Haas P, van Deutekom H, Roring S, Bifani P, Kurepina N, Kreiswirth B, Sola C, Rastogi N, Vatin V, Gutierrez MC, Fauville M, Niemann S, Skuce R, Kremer K, Locht C, van Soolingen D. 2006. Proposal for standardization of optimized mycobacterial interspersed repetitive unit-variable-number tandem repeat typing of Mycobacterium tuberculosis. J Clin Microbiol 44:4498–4510. 154. Supply P, Mazars E, Lesjean S, Vincent V, Gicquel B, Locht C. 2000. Variable human minisatellite-like regions in the Mycobacterium tuberculosis genome. Mol Microbiol 36:762–771. 155. Mazars E, Lesjean S, Banuls AL, Gilbert M, Vincent V, Gicquel B, Tibayrenc M, Locht C, Supply P. 2001. High-resolution minisatellite-based typing as a portable approach to global analysis of Mycobacterium tuberculosis molecular epidemiology. Proc Natl Acad Sci USA 98:1901–1906. 156. Allix-Beguec C, Fauville-Dufaux M, Supply P. 2008. Three-year population-based evaluation of standardized mycobacterial interspersed repetitive-unit-variable-number tandem-repeat typing of Mycobacterium tuberculosis. J Clin Microbiol 46:1398–1406. 157. Oelemann MC, Diel R, Vatin V, Haas W, Rusch-Gerdes S, Locht C, Niemann S, Supply P. 2007. Assessment of an optimized mycobacterial interspersed repetitive- unitvariable-number tandem-repeat typing system combined with spoligotyping for population-based molecular epidemiology studies of tuberculosis. J Clin Microbiol 45:691– 697. 158. Pitondo-Silva A, Santos AC, Jolley KA, Leite CQ, Darini AL. 2013. Comparison of three molecular typing methods to assess genetic diversity for Mycobacterium tuberculosis. J Microbiol Methods 93:42–48. 159. Alvarez J, Garcia IG, Aranaz A, Bezos J, Romero B, de Juan L, Mateos A, Gomez-Mampaso E, Dominguez L. 2008. Genetic diversity of Mycobacterium avium isolates recovered from clinical samples and from the environment: molecular characterization for diagnostic purposes. J Clin Microbiol 46:1246–1251. 160. Conger NG, O’Connell RJ, Laurel VL, Olivier KN, Graviss EA, Williams-Bouyer N, Zhang Y, Brown-Elliott BA, Wallace RJ Jr. 2004. Mycobacterium simae outbreak associated with a hospital water supply. Infect Control Hosp Epidemiol 25:1050–1055. 161. Doig C, Muckersie L, Watt B, Forbes KJ. 2002. Molecular epidemiology of Mycobacterium malmoense infections in Scotland. J Clin Microbiol 40:1103–1105. 162. Garriga X, Cortes P, Rodriguez P, March F, Prats G, Coll P. 2000. Comparison of IS1245 restriction fragment length polymorphism and pulsed-field gel electrophoresis for typing clinical isolates of Mycobacterium avium subsp avium. Int J Tuberc Lung Dis 4:463–472. 163. Sevilla I, Li L, Amonsin A, Garrido JM, Geijo MV, Kapur V, Juste RA. 2008. Comparative analysis of Mycobacterium avium subsp. paratuberculosis isolates from cattle, sheep and goats by short sequence repeat and pulsed-field gel electrophoresis typing. BMC Microbiol 8:204. 164. Pfaller SL, Aronson TW, Holtzman AE, Covert TC. 2007. Amplified fragment length polymorphism analysis of Mycobacterium avium complex isolates recovered from southern California. J Med Microbiol 56:1152–1160. 165. Turenne CY, Wallace R Jr, Behr MA. 2007. Mycobacterium avium in the postgenomic era. Clin Microbiol Rev 20:205–229. 166. Griffith DE, Brown-Elliott BA, Langsjoen B, Zhang Y, Pan X, Girard W, Nelson K, Caccitolo J, Alvarez J, Shepherd S, Wilson R, Graviss EA, Wallace RJ Jr. 2006. Clinical and molecular analysis of macrolide resistance

167.

168.

169.

170.

171.

172.

173.

174.

175.

176.

177.

178.

179.

in Mycobacterium avium complex lung disease. Am J Respir Crit Care Med 174:928–934. Holmes GP, Bond GB, Fader RC, Fulcher SF. 2002. A Cluster of cases of Mycobacterium szulgai keratitis that occurred after laser-assisted in situ keratomileusis. Clin Infect Dis 34:1039–1046. Picardeau M, Prod’Hom G, Raskine L, LePennec MP, Vincent V. 1997. Genotypic characterization of five subspecies of Mycobacterium kansasii. J Clin Microbiol 35:25– 32. Wallace RJ Jr, Zhang Y, Brown BA, Dawson D, Murphy DT, Wilson R, Griffith DE. 1998. Polyclonal Mycobacterium avium complex infections in patients with nodular bronchiectasis. Am J Respir Crit Care Med 158:1235– 1244. Wallace RJ, Zhang Y, Brown-Elliott BA, Yakrus MA, Wilson RW, Mann L, Couch L, Girard WM, Griffith DE. 2002. Repeat positive cultures in Mycobacterium intracellulare lung disease after macrolide therapy represent new infections in patients with nodular bronchiectasis. J Infect Dis 186:266–273. Zhang Y, Mann LB, Wilson RW, Brown-Elliott BA, Vincent V, Iinuma Y, Wallace RJ Jr. 2004. Molecular analysis of Mycobacterium kansasii isolates from the United States. J Clin Microbiol 42:119–125. Ashworth M, Horan KL, Freeman R, Oren E, Narita M, Cangelosi GA. 2008. Use of PCR-based Mycobacterium tuberculosis genotyping to prioritize tuberculosis outbreak control activities. J Clin Microbiol 46:856–862. Cangelosi GA, Freeman RJ, Lewis KN, Livingston-Rosanoff D, Shah KS, Milan SJ, Goldberg SV. 2004. Evaluation of a high-throughput repetitive-sequence-based PCR system for DNA fingerprinting of Mycobacterium tuberculosis and Mycobacterium avium complex strains. J Clin Microbiol 42:2685–2693. Horan KL, Freeman R, Weigel K, Semret M, Pfaller S, Covert TC, van Soolingen D, Leao SC, Behr MA, Cangelosi GA. 2006. Isolation of the genome sequence strain Mycobacterium avium 104 from multiple patients over a 17-year period. J Clin Microbiol 44:783–789. Inagaki T, Nishimori K, Yagi T, Ichikawa K, Moriyama M, Nakagawa T, Shibayama T, Uchiya K, Nikai T, Ogawa K. 2009. Comparison of a variable-number tandem-repeat (VNTR) method for typing Mycobacterium avium with mycobacterial interspersed repetitive-unitVNTR and IS1245 restriction fragment length polymorphism typing. J Clin Microbiol 47:2156–2164. Mobius P, Luyven G, Hotzel H, Kohler H. 2008. High genetic diversity among Mycobacterium avium subsp. paratuberculosis strains from German cattle herds shown by combination of IS900 restriction fragment length polymorphism analysis and mycobacterial interspersed repetitive unit-variable-number tandem-repeat typing. J Clin Microbiol 46:972–981. Thibault VC, Grayon M, Boschiroli ML, Hubbans C, Overduin P, Stevenson K, Gutierrez MC, Supply P, Biet F. 2007. New variable-number tandem-repeat markers for typing Mycobacterium avium subsp. paratuberculosis and M. avium strains: comparison with IS900 and IS1245 restriction fragment length polymorphism typing. J Clin Microbiol 45:2404–2410. Wallace RJ Jr, Iakhiaeva E, Williams MD, Brown-Elliott BA, Vasireddy S, Vasireddy R, Lande L, Peterson DD, Sawicki J, Kwait R, Tichenor WS, Turenne C, Falkinham JO III. 2013. Absence of Mycobacterium intracellulare and presence of Mycobacterium chimaera in household water and biofilm samples of patients in the United States with Mycobacterium avium complex respiratory disease. J Clin Microbiol 51:1747–1752. van Soolingen D, Bauer J, Ritacco V, Leao SC, Pavlik I, Vincent V, Rastogi N, Gori A, Bodmer T, Garzelli

594 n

180.

181.

182.

183.

184.

185.

186.

187.

188.

189.

190.

BACTERIOLOGY

C, Garcia MJ. 1998. IS1245 restriction fragment length polymorphism typing of Mycobacterium avium isolates: proposal for standardization. J Clin Microbiol 36:3051– 3054. Mijs W, de Haas P, Rossau R, Van der Laan T, Rigouts L, Portaels F, van Soolingen D. 2002. Molecular evidence to support a proposal to reserve the designation Mycobacterium avium subsp. avium for bird-type isolates and ‘M. avium subsp. hominissuis’ for the human/porcine type of M. avium. Int J Syst Evol Microbiol 52:1505–1518. Turenne CY, Collins DM, Alexander DC, Behr MA. 2008. Mycobacterium avium subsp. paratuberculosis and M. avium subsp. avium are independently evolved pathogenic clones of a much broader group of M. avium organisms. J Bacteriol 190:2479–2487. Clinical and Laboratory Standards Institute. 2009. Susceptibility Testing of Mycobacteria, Nocardiae, and other Aerobic Actinomycetes; Approved Standard—2nd Edition. M24A2. CLSI, Wayne, PA. Brown-Elliott BA, Iakhiaeva E, Griffith DE, Woods GL, Stout JE, Wolfe CR, Turenne CY, Wallace RJ Jr. 2013. In vitro activity of amikacin against isolates of Mycobacterium avium complex with proposed MIC breakpoints and finding of a 16S rRNA gene mutation in treated isolates. J Clin Microbiol 51:3389–3394. Abuali MM, Katariwala R, LaBombardi VJ. 2012. A comparison of the Sensititre(R) MYCOTB panel and the agar proportion method for the susceptibility testing of Mycobacterium tuberculosis. Eur J Clin Microbiol Infect Dis 31:835–839. Hall L, Jude KP, Clark SL, Dionne K, Merson R, Boyer A, Parrish NM, Wengenack NL. 2012. Evaluation of the Sensititre MycoTB plate for susceptibility testing of the Mycobacterium tuberculosis complex against first- and second-line agents. J Clin Microbiol 50:3732–3734. Almeida Da Silva PE, Palomino JC. 2011. Molecular basis and mechanisms of drug resistance in Mycobacterium tuberculosis: classical and new drugs. J Antimicrob Chemother 66:1417–1430. Campbell PJ, Morlock GP, Sikes RD, Dalton TL, Metchock B, Starks AM, Hooks DP, Cowan LS, Plikaytis BB, Posey JE. 2011. Molecular detection of mutations associated with first- and second-line drug resistance compared with conventional drug susceptibility testing of Mycobacterium tuberculosis. Antimicrob Agents Chemother 55:2032–2041. Thorel MF, Krichevsky M, Levy-Frebault VV. 1990. Numerical taxonomy of mycobactin-dependent mycobacteria, emended description of Mycobacterium avium, and description of Mycobacterium avium subsp. avium subsp. nov., Mycobacterium avium subsp. paratuberculosis subsp. nov., and Mycobacterium avium subsp. silvaticum subsp. nov. Int J Syst Bacteriol 40:254–260. Bang D, Herlin T, Stegger M, Andersen AB, Torkko P, Tortoli E, Thomsen VO. 2008. Mycobacterium arosiense sp. nov., a slowly growing, scotochromogenic species causing osteomyelitis in an immunocompromised child. Int J Syst Evol Microbiol 58:2398–2402. Tortoli E, Rindi L, Garcia MJ, Chiaradonna P, Dei R, Garzelli C, Kroppenstedt RM, Lari N, Mattei R, Mariottini A, Mazzarelli G, Murcia MI, Nanetti A, Piccoli P, Scarparo C. 2004. Proposal to elevate the genetic variant MAC-A, included in the Mycobacterium avium complex, to species rank as Mycobacterium chimaera sp. nov. Int J Syst Evol Microbiol 54:1277–1285.

191. Murcia MI, Tortoli E, Menendez MC, Palenque E, Garcia MJ. 2006. Mycobacterium colombiense sp. nov., a novel member of the Mycobacterium avium complex and description of MAC-X as a new ITS genetic variant. Int J Syst Evol Microbiol 56:2049–2054. 192. van Ingen J, Boeree MJ, Kosters K, Wieland A, Tortoli E, Dekhuijzen PN, van Soolingen D. 2009. Proposal to elevate Mycobacterium avium complex ITS sequevar MAC-Q to Mycobacterium vulneris sp. nov. Int J Syst Evol Microbiol 59:2277–2282. 193. Reischl U, Emler S, Horak Z, Kaustova J, Kroppenstedt RM, Lehn N, Naumann L. 1998. Mycobacterium bohemicum sp. nov., a new slow-growing scotochromogenic mycobacterium. Int J Syst Bacteriol 48 Pt 4:1349–1355. 194. Bull TJ, Shanson DC, Archard LC, Yates MD, Hamid ME, Minnikin DE. 1995. A new group (type 3) of Mycobacterium celatum isolated from AIDS patients in the London area. Int J Syst Bacteriol 45:861–862. 195. Butler WR, O’Connor SP, Yakrus MA, Smithwick RW, Plikaytis BB, Moss CW, Floyd MM, Woodley CL, Kilburn JO, Vadney FS, Gross WM. 1993. Mycobacterium celatum sp. nov. Int J Syst Bacteriol 43:539–548. 196. Reischl U, Feldmann K, Naumann L, Gaugler BJ, Ninet B, Hirschel B, Emler S. 1998. 16S rRNA sequence diversity in Mycobacterium celatum strains caused by presence of two different copies of 16S rRNA gene. J Clin Microbiol 36:1761–1764. 197. Bottger EC, Hirschel B, Coyle MB. 1993. Mycobacterium genavense sp. nov. Int J Syst Bacteriol 43:841–843. 198. Haas WH, Butler WR, Kirschner P, Plikaytis BB, Coyle MB, Amthor B, Steigerwalt AG, Brenner DJ, Salfinger M, Crawford JT, Bottger EC, Bremer HJ. 1997. A new agent of mycobacterial lymphadenitis in children: Mycobacterium heidelbergense sp. nov. J Clin Microbiol 35:3203– 3209. 199. Springer B, Kirschner P, Rost-Meyer G, Schroder KH, Kroppenstedt RM, Bottger EC. 1993. Mycobacterium interjectum, a new species isolated from a patient with chronic lymphadenitis. J Clin Microbiol 31:3083–3089. 200. Meier A, Kirschner P, Schroder KH, Wolters J, Kroppenstedt RM, Bottger EC. 1993. Mycobacterium intermedium sp. nov. Int J Syst Bacteriol 43:204–209. 201. Iwamoto T, Fujiyama R, Yoshida S, Wada T, Shirai C, Kawakami Y. 2009. Population structure dynamics of Mycobacterium tuberculosis Beijing strains during past decades in Japan. J Clin Microbiol 47:3340–3343. 202. Taillard C, Greub G, Weber R, Pfyffer GE, Bodmer T, Zimmerli S, Frei R, Bassetti S, Rohner P, Piffaretti JC, Bernasconi E, Bille J, Telenti A, Prod’hom G. 2003. Clinical implications of Mycobacterium kansasii species heterogeneity: Swiss National Survey. J Clin Microbiol 41:1240–1244. 203. Tonjum T, Welty DB, Jantzen E, Small PL. 1998. Differentiation of Mycobacterium ulcerans, M. marinum, and M. haemophilum: mapping of their relationships to M. tuberculosis by fatty acid profile analysis, DNA-DNA hybridization, and 16S rRNA gene sequence analysis. J Clin Microbiol 36:918–925. 204. Kent PT, Kubica GP. 1985. Public Health Mycobacteriology. A Guide for the Level III Laboratory. Centers for Disease Control and Prevention U.S. Department of Health and Human Services, Atlanta, GA. 205. Richter E, Greinert U, Kirsten D, Rusch-Gerdes S, Schluter C, Duchrow M, Galle J, Magnussen H, Schlaak M, Flad HD, Gerdes J. 1996. Assessment of mycobacterial DNA in cells and tissues of mycobacterial and sarcoid lesions. Am J Respir Crit Care Med 153:375–380.

Mycobacterium: Clinical and Laboratory Characteristics of Rapidly Growing Mycobacteria BARBARA A. BROWN-ELLIOTT AND RICHARD J. WALLACE, JR.

32 abscessus subsp. massiliense is composed of several different genotypes, which are also related to colony morphology (13). Thus, for ease of reading in this chapter, we use the terminology “M. abscessus subsp. massiliense” to refer to the former species M. massiliense. Two newly described species, M. bacteremicum and “M. franklinii,” were described in 2011, although the latter species has not yet been validated (4, 14). The second recently validated species, M. litorale, was described from soil samples in China in 2012 but has not been recovered from clinical samples (3). A third newly described species, M. iranicum, has been reported as a pathogen in the United States, Europe, and Asia and has been isolated from respiratory, wound, and cerebrospinal fluid cultures (5). There are currently six major groups or complexes of RGM, based on pigmentation and genetic relatedness (Table 1). The first group of nonpigmented pathogenic species includes 10 species, some of which once belonged to the M. fortuitum group (Table 1) (1, 15–19). The second group of nonpigmented RGM is the M. chelonae/M. abscessus group listed in Table 1 (1, 7, 20–23). A previously described species, M. salmoniphilum, has been revived and also considered to be related to this group. Although this species has been recovered from disseminated disease of salmon and trout, it has not been recovered from humans (24). “M. franklinii,” the currently nonvalidated species, has been described for multiple patients with sinopulmonary disease, primarily from the northeastern United States (14). Six isolates from extrapulmonary sources have also been described. The organism is closely related to the M. chelonae/M. abscessus group. A third nonpigmented group, the M. mucogenicum group, currently includes three species, as noted in Table 1 (1, 20, 25). The fourth group, the M. smegmatis group, is currently composed of the two late-pigmented species, i.e., M. smegmatis (formerly known as M. smegmatis sensu stricto) and M. goodii (1, 16, 26). The fifth group of RGM includes the early-pigmented species, which traditionally have been difficult to identify by conventional (phenotypic) laboratory methods. The only proven pathogens in this group are M. neoaurum and M. bacteremicum (Table 1), which have been associated with mycobacteremia in patients with catheter infections, and M. iranicum, which has been recovered from five respiratory

TAXONOMY AND DESCRIPTION OF THE AGENTS The rapidly growing mycobacteria (RGM) are generally defined as nontuberculous species that grow on laboratory media within 7 days (1). RGM contain long-chain fatty acids, known as mycolic acids, that can be quantitated using chromatographic techniques, such as high-performance liquid chromatography (HPLC). Prior to the molecular era, HPLC was used for identification of species of the RGM in most major reference laboratories. However, this method has been replaced in most laboratories by more definitive molecular identification methods for more accurate species identification (2). Currently, there are more than 140 species of nontuberculous mycobacteria (NTM), among which approximately 70 are species of RGM. More than half of the RGM species have been described since the early 1990s. Since 2010, only three new validated species, Mycobacterium litorale, M. bacteremicum, and M. iranicum, have been added to the list (3–5). Although the species M. algericum was described as an RGM, only microcolonies were evident at 1 week. Moreover, this organism is most closely related to the slowly growing mycobacteria of the M. terrae complex and thus, by definition, should not be considered an RGM (6). Additionally, most recently, the species M. abscessus was subdivided into two subspecies. The former species M. abscessus is now designated M. abscessus subsp. abscessus, and two previously described species, M. massiliense and M. bolletii, have been combined and proposed to form another subspecies of M. abscessus, M. abscessus subsp. bolletii (7, 8). The latter designation is controversial since the recent discovery that no functional erm gene is present in isolates previously named M. massiliense, in contrast to the case for isolates previously called M. bolletii, with a functional erm gene conferring inducible macrolide resistance. Moreover, the 2012 descriptions of the genomic sequences of M. abscessus subsp. bolletii (previously M. bolletii) and M. abscessus subsp. massiliense (previously M. massiliense) may help to resolve some of the questions surrounding the species and provide information to better discriminate between the subspecies (9, 10). Interestingly, Choi et al. noted two rRNA gene operons in the species, which is a distinctly different finding from previous reports of a single rRNA gene operon in M. abscessus (11, 12). Moreover, recent studies showed that M.

doi:10.1128/9781555817381.ch32

595

596 n

BACTERIOLOGY TABLE 1

Six major groups of RGM

Group or taxon

Species within group or taxon

Mycobacterium fortuitum group ............................... M. fortuitum, M. peregrinum, M. senegalense, M. setense, M. septicum,a M. porcinum,a M. houstonense,a M. boenickei,a M. brisbanense,a M. neworleansensea M. chelonae/M. abscessus group ............................. M. chelonae, M. immunogenum, M. abscessus subsp. abscessus (formerly M. abscessus), M. abscessus subsp. bolletii (formerly M. massiliense and M. bolletii), M. salmoniphilum, “M. franklinii”b M. mucogenicum group ........................................... M. mucogenicum, M. aubagnense, M. phocaicum M. smegmatis group ................................................ M. smegmatis (formerly M. smegmatis sensu stricto), M. goodii Early pigmented RGM ........................................... M. neoaurum, M. canariasense, M. cosmeticum, M. monacense, M. bacteremicum M. mageritense/M. wolinskyi group ......................... M. mageritense, M. wolinskyi a

Includes the former third-biovariant complex. “M. franklinii” is a nonvalidated species.

b

samples, a hand wound, and two cerebrospinal fluid samples (4, 5, 27). Several newly described early-pigmented species, including M. canariasense, M. cosmeticum, and M. monacense, are also clinically significant (28–30). There are a number of previously listed environmental (nonpathogenic) species as well (1, 17, 28, 31–33). Current studies based on DNA sequence analysis suggest the presence of a sixth, nonpigmented pathogenic group, composed of M. mageritense and M. wolinskyi (2, 21, 22). The introduction of the new species noted above within the RGM emphasizes the importance of molecular identification to the species level and questions the meaningfulness of the current “group” classifications, especially for the M. fortuitum group. However, because previous data (prior to the reliance on genetic testing) and publications use this “group” nomenclature, these designations are retained in this chapter for ease of discussion (1). As more information is obtained using advanced molecular technology and more laboratories adopt this newer methodology, the group designations may become even more obsolete.

CLINICAL SIGNIFICANCE The RGM are opportunistic pathogens that produce disease in a variety of clinical settings. The three major clinically important species of RGM, responsible for approximately 80% of mycobacterial disease in humans, are the M. fortuitum group (including M. porcinum), M. chelonae, and M. abscessus (19, 34, 35). Other potentially pathogenic and clinically significant RGM species are included in Table 2 (1, 2, 15–17, 20–24, 26, 28, 30, 32, 33, 36–39). RGM are presumed to be common in the environment and have been identified most often in tap water when associated with outbreaks of catheter sepsis in bone marrow transplants, wound infections, and associated pseudo-outbreaks of disease (1, 35). The specific reservoir for M. abscessus chronic lung infections has yet to be identified.

Community-Acquired Skin and Soft Tissue Infections The most common infection seen with the RGM is a posttraumatic-wound infection. The patients are generally healthy, and drug-induced immune suppression shows a minimal increase in risk for this type of infection. The

M. fortuitum group accounts for approximately 60% of cases of localized cutaneous infections, but any of the more than 30 pathogenic RGM species listed in Table 2 can cause disease (1, 15, 33, 40, 41). Traumatic wound infections, especially open fractures, often involve species within the former Mycobacterium fortuitum third-biovariant complex (i.e., M. porcinum, and M. houstonense) (Table 1) (1, 40). More than 75% of the infections reported in a series of 85 isolates of the M. fortuitum third biovariant from the United States and the Queensland, Australia, state laboratory were associated with skin, soft tissue, or bone infections (1). The majority of infections occurred 4 to 6 weeks following puncture wounds or open fractures. Metal puncture wounds (48%) and injuries from motor vehicle accidents (26%) were the most common antecedent injuries, and approximately 40% of the injury sites involved the foot or leg. Stepping on a nail was the most frequently related scenario. None of the isolates in this series were studied by molecular techniques that would identify them as one of the species within the former M. fortuitum third-biovariant complex. A 2011 report of an outbreak from 2003 until 2010 in a large university medical center in Texas, involving M. porcinum, included not only multiple respiratory isolates but also environmental water and ice isolates, along with clinical isolates from localized abscesses (spinal and paraspinal sites, finger, neck, and breast), peritoneal fluid, blood, and/or Port-a-Cath sites (35). Sporadic cases of localized wound infections following medical or surgical procedures, including needle injections, can occur with M. chelonae but are less common than those with M. fortuitum. In late 2011 and early 2012, an outbreak of 19 cases of M. chelonae associated with tattoo ink were reported in New York (42). This was the largest documented outbreak of this type involving M. chelonae. The clinical picture of posttraumatic-wound infection ranges from localized cellulitis or abscesses to osteomyelitis (1). A 2006 report from a major U.S. clinical referral center for patients from Minnesota, Wisconsin, Iowa, and South Dakota characterized 63 HIVnegative patients with RGM infections involving M. abscessus or M. chelonae (71%) or involving the M. fortuitum group (29%). Moreover, patients with M. chelonae or M. abscessus usually had multiple (disseminated) cutaneous

32. Rapidly Growing Mycobacteria n TABLE 2

597

Currently recognized species of RGMl

Pigmentation

Unique phenotype

Unique hsp65 PRA pattern

Unique (complete) 16S rRNA gene sequence

Yes Yes Yes

No No No

No Yes Yes

Yes Yes Yes

Yes Yes Yes

Less common human pathogens (>10 clinical isolates or cases) M. abscessus subsp. bolletiib M. abscessus subsp. massilienseb M. bacteremicum M. boenickei M. canariasense M. cosmeticum “M. franklinii”c M. goodii M. houstonense M. immunogenum M. mageritense M. mucogenicum M. neoaurum M. peregrinum M. porcinum M. senegalense M. smegmatis

Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes Yes

No No Yes No No Yes No Yes No No No No Yes No No No Yes

No No No No No Yes No No No No Yes Yes No No No No No

No No Yes Yes Yes Yes Yes Yes No Yes Yes Yes Yes Yes Yes Yes Yes

No No Yes Yes Yes Yes No Yes Nod Yes Yes Yes Yes Noe Yes Yes Yes

Rare human pathogens (80% of cases of pulmonary disease due to RGM (34). Similarities exist between patients with MAC and those with M. abscessus, such that a common pathogenicity or host susceptibility factor may be involved (34). Multiple cultures of M. abscessus from respiratory samples are usually associated with significant pulmonary disease.

Cystic Fibrosis M. abscessus subsp. abscessus and M. abscessus subsp. massiliense have been isolated with increasing frequency from the respiratory tracts of patients with cystic fibrosis (CF) (1, 57–62). A 2012 study concluded that CF patients with M. abscessus were younger and had more severe disease and more frequent intravenous antimicrobial treatment (62). M. abscessus is the second most common species of nontuberculous mycobacteria recovered from CF patients (after the MAC), and it may be the most common species associated with clinical disease in this setting (59). Patients with CF also have bronchiectasis, in addition to chronic, recur-

Hypersensitivity pneumonitis among metal grinders working with contaminated metalworking fluids in industrial plants has been associated with a newly described species of RGM, M. immunogenum (1, 34, 65). Multiple pseudooutbreaks associated with this species have been reported, resulting from contaminated automated bronchoscope cleaning machines and metalworking fluids. This species is able to grow and remain viable in degraded metalworking fluid and is resistant to the routine biocides used for disinfection of metalworking fluids (1, 65). However, as yet, this species has not been reported from open lung biopsy specimens from these patients.

Central Nervous System Disease Central nervous system disease involving the RGM is rare, but morbidity and mortality are high. Most of the reported cases have been associated with M. fortuitum (1, 66, 67).

Corneal Infections (Keratitis) The number of RGM recovered from ocular infections has been increasing over the last 20 years. A retrospective review of cases of NTM keratitis from 1982 to 1997 at an eye institute in Florida showed that 19 of 24 cases were due to RGM (68). A recent study which identified 113 ophthalmic isolates by molecular methods showed that the most common RGM were M. chelonae (45%), M. abscessus (42%), and the M. fortuitum group (8%) (69).

600 n BACTERIOLOGY

Since the early 1990s, other descriptions of epidemic and sporadic ocular infections associated with the RGM have been published, including postkeratoplasty infections and infections following laser in situ keratomileusis (LASIK) surgery (1, 69).

Otitis Media The most common NTM ear disease is chronic otitis media due to M. abscessus. In a 1988 outbreak of 17 cases of otitis media in two ear-nose-throat clinics, patients presented with chronic ear drainage with a perforated tympanic membrane and a prior tympanostomy tube (1). In another series, 20 of 21 cases of sporadic chronic otitis media (some with associated mastoiditis) were due to M. abscessus infection following ear tube placement. Approximately one-half of the isolates from these cases were aminoglycoside resistant, resulting from the long-term use of aminoglycoside ear drops (1). Sporadic cases of this disease continue to be seen. The key to the disease appears to be the presence of the foreign body (tympanostomy tube). Recently, otitis media and otomastoiditis caused by M. abscessus subsp. massiliense were described for middle-aged adults with underlying otologic disease. These patients were treated successfully with clarithromycin-containing regimens for up to 9 months (70).

Health Care-Associated Infections Health care-associated disease with RGM has been reported most commonly with M. fortuitum, M. chelonae, M. abscessus, and M. mucogenicum, although any species may be involved. Most infections occur following contamination with tap water (1, 40, 71, 72). Types of infections include postsurgical wound infections, catheter sepsis, infections following hemodialysis, postinjection abscesses, vaccine-related outbreaks, and otitis media following tympanostomy tube replacement (1, 73, 74). These have been seen as both sporadic cases and localized outbreaks. Recent outbreaks have involved cosmetic procedures, such as liposuction, liposculpture, acupuncture, and mesotherapy, a procedure comprising multiple subcutaneous injections of pharmaceutical or homeopathic medications for cosmetic purposes (41, 73–80). In addition to true outbreaks of infection, numerous health care-associated pseudo-outbreaks have been described. Contaminated or malfunctioning bronchoscopes, automated endoscope cleaning machines, and contaminated laboratory reagents and ice have been implicated (1, 73, 81–83).

commonly associated with augmentation mammaplasty and coronary artery bypass surgery, and multiple disease outbreaks occurred (1, 87). Infections following these types of surgery are now less common, although a cluster of 12 cases of post-augmentation mammaplasty surgical site infections due to M. fortuitum and M. porcinum was reported between 2002 and 2004 in Brazil (87). More often, however, these types of infections have been replaced by infections following other types of cosmetic surgeries, such as liposuction, and other types of prosthetic surgeries, such as knee replacements. In a 1989 report (15), approximately 80% of RGM wound isolates related to cardiac surgery were from seven southern coastal states, including Texas, Louisiana, Georgia, Maryland, Alabama, Florida, and South Carolina. A second report, published in the same year, showed that 92% of 37 identified cases of surgical wound infection following augmentation mammaplasty were also from patients in southern coastal states, with the majority located in Texas, Florida, and North Carolina, suggesting that the disease risk was highest in the southeastern United States (1). Recovery of both subspecies of M. abscessus has been reported from outbreaks of infections associated with laparoscopic surgeries and cosmetic surgeries in Brazil, the Dominican Republic, and Korea (76, 78). These and other recent reports suggest that although few studies have identified these newer subspecies in invasive infections, they have been misclassified in previous studies (88).

Prosthetic Device Infections Infections following insertion of prosthetic devices, including prosthetic heart valves, artificial knees and hips, lens implants, and metal rods inserted into the vertebrae to stabilize bones following fractures, have also been described (1, 52). Again, M. fortuitum is the most common pathogen, but any of the pathogenic RGM, including the M. smegmatis group, can be associated with this type of infection (1).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS Details of standard methods are included in chapter 30. Transport of species is accomplished by using leakproof containers and proper safety protocols. Specimens for detection of RGM should be delivered to the laboratory in a timely manner following appropriate shipping and handling regulations for shipping biological or infectious materials (89).

Catheter-Associated Infections Central catheter-associated infections are the most common health care-associated infections due to the RGM (1, 39, 85). They are the most common cause of RGM bacteremia, but the disease may also present as local wound drainage as part of an exit site or tunnel infection (1, 85). Other types of catheters can also become infected, including peritoneal catheters, ventriculo-peritoneal shunts, and shunts for hemodialysis (1). The most common species are M. mucogenicum, M. fortuitum, M. abscessus, and the newly described species M. bacteremicum. An outbreak of M. phocaicum and M. mucogenicum was described for five patients with central venous catheters in an oncology unit in a Texas hospital (86). This outbreak represents the first report of clinical isolates of M. phocaicum in a hospital in the United States.

Disinfectants The RGM are generally resistant to the activity of biocides, such as organomercurials, chlorine, 2% formaldehyde, and alkaline glutaraldehyde, all of which are commonly used disinfectants (1). A report of the contamination of benzalkonium chloride, a widely used antiseptic compound, with M. abscessus and of a resulting outbreak of this species in several patients following steroid injections after skin disinfection emphasizes the limitations of disinfectants against the RGM (90).

DIRECT EXAMINATION Microscopy

Surgical Wound Infections Surgical wound infections due to RGM are a well-recognized clinical entity. In the 1970s and 1980s, these were most

The use of mycobacterial smears is a rapid and reasonably sensitive step in the diagnosis of RGM disease. Gram stains of colonies showing faintly staining, “ghost-like” beaded

32. Rapidly Growing Mycobacteria n 601

Gram-positive bacilli are often helpful in establishing a diagnosis of mycobacteriosis. Ziehl-Neelsen or Kinyoun staining may also be useful. However, the use of smears alone is not sufficient for species identification. A large study found that NTM, including RGM, are likely to be detected by fluorochrome staining of specimens, especially those from patients at low risk for AIDS in areas where lung disease is endemic (89, 91). Further details of the staining procedures are found in chapter 30.

Nucleic Acid Detection Currently, there are no commercial nucleic acid detection systems available for direct detection of NTM. An indirect method could be identifying an acid-fast bacillus smearpositive sample that is negative by a nucleic acid amplification test specific for Mycobacterium tuberculosis, but this does not prove the presence of RGM compared to the slowly growing NTM.

ISOLATION PROCEDURES Primary isolation of RGM optimally requires culture at 28 to 30°C rather than 35°C, especially for recovery of M. chelonae and M. immunogenum (1). Direct examination and isolation procedures are detailed in chapter 30. Recovery of RGM from routine blood culture systems does not appear to be problematic. However, no large comparative studies of commercial systems have been performed to date.

IDENTIFICATION Biochemical Testing As previously stated, the RGM are defined as NTM that grow within 7 days (most species grow within 3 to 4 days) (1). Until the advent of more modern molecular techniques, traditional laboratory identification of the RGM was based primarily upon growth rate, pigmentation, colonial morphology, and a select battery of biochemical tests (1). These standard tests included tests of arylsulfatase production, tolerance to 5% NaCl, nitrate reductase activity, and iron uptake. All members of the M. fortuitum group and M. chelonae/M. abscessus group exhibit a strongly positive arylsulfatase reaction at 3 days. The M. smegmatis group (M. smegmatis and M. goodii) and M. wolinskyi have similar growth rates but do not exhibit arylsulfatase activity at 3 days (1, 26). Approximately 95% of the isolates of M. smegmatis (sensu stricto) and 80% of M. goodii isolates develop a late (7 to 10 days) yellow-orange pigmentation (1, 26). The current proposal for the clinical laboratory is that biochemical testing of the RGM should be replaced by molecular methods. Moreover, biochemical testing should be performed only when describing a new species as part of a polyphasic identification algorithm.

Supplemental Biochemical Testing: Carbohydrate Utilization The supplementation of standard biochemical tests with carbohydrate utilization has allowed more complete and accurate laboratory identification of established species and discrimination of some (but not all) new species (1). Identification to the species level and susceptibility testing (see chapter 76) should be performed on isolates of RGM considered to be clinically significant.

However, as previously stated, molecular testing is the only definitive means of identifying the RGM species, and laboratories should proceed cautiously in identifying these species by biochemical testing alone (92, 93).

Antimicrobial Susceptibility Tests for Taxonomic Identification As discussed above, other adjunctive nonmolecular tests, including antimicrobial susceptibility tests, have also been utilized for identification of the RGM (1, 93). They are less commonly used currently, with the advent of molecular techniques, but can provide species confirmation in conjunction with molecular results. As a screening tool, isolates of the M. fortuitum group and the M. chelonae/M. abscessus group can be differentiated by the use of a polymyxin B (300 IU) disk diffusion method. Generally, isolates of the M. fortuitum group exhibit a partial or clear zone of inhibition (≥10 mm) around the polymyxin disk, whereas isolates of the M. chelonae/M. abscessus group show no zone of inhibition (1). Moreover, M. chelonae and M. abscessus also have different antimicrobial susceptibility patterns. One major difference between the two species is resistance to cefoxitin. By agar disk diffusion, M. chelonae shows complete resistance to cefoxitin, with no partial or complete zones of inhibition, in contrast to the partial or complete zones seen with M. abscessus. Isolates of M. chelonae generally have cefoxitin MICs of ≥256 μg/ml, whereas the modal MIC for isolates of M. abscessus is 32 to 64 μg/ml (1). Furthermore, recent studies have shown that isolates of M. abscessus subsp. abscessus, but not M. chelonae or M. abscessus subsp. massiliense, have an inducible erm gene similar to the gene in M. fortuitum, which conveys macrolide resistance (94, 95). Isolates of M. abscessus also have lower amikacin MICs and are resistant to tobramycin, whereas tobramycin is more active than amikacin with M. chelonae. Additionally, isolates of M. chelonae are more susceptible in vitro to some of the newer antibiotics, including linezolid and moxifloxacin, than are isolates of M. abscessus (92).

HPLC Identification HPLC analysis of the mycobacterial cell wall mycolic acid content is routinely used in large reference or state health department laboratories to identify slowly growing isolates of NTM, but it has been problematic with RGM (1, 96). HPLC can be helpful for placing RGM isolates into groups or complexes, but it is not specific enough to identify most species with a high degree of accuracy.

Molecular Identification Methods Nucleic Acid Probes The INNO-LiPA multiplex probe assay (Innogenetics, Ghent, Belgium) is based on the principle of reverse hybridization (97). Although the assay has not received U.S. Food and Drug Administration clearance, it can identify both rapidly and slowly growing mycobacterial species. Biotinylated DNA obtained by PCR amplification of the 16S-23S internal transcribed spacer (ITS) is hybridized with specific oligonucleotide probes immobilized as parallel lines on membrane strips. The main advantage of this system is that a large variety of species may be identified by a single assay, without the need to select an appropriate probe. One limitation of the assay is the cross-reactivity that may be detected with species of the M. fortuitum group and several species, such as M. thermoresistibile, M. agri, and M. alvei, that are rarely found in clinical samples (1, 97). Additionally, it

602 n

BACTERIOLOGY

failed to differentiate isolates of closely related species, such as M. chelonae and M. abscessus (98). Since the original studies with the INNO-LiPA assay, however, the system has been improved to include additional probes for the M. fortuitum-peregrinum complex and M. smegmatis. A similar commercial PCR method which targets the 23S rRNA gene, the GenoType Mycobacterium assay (Hain Lifescience, GmbH, Nehren, Germany), provides probes for simultaneous identification of M. chelonae and specific probes for M. peregrinum, M. fortuitum, and M. phlei. These two systems are widely used in Europe for NTM identification (99, 100).

Sequence Analysis Nucleic acid sequence analysis has been performed for the identification of mycobacteria for several years. This identification tool has been useful for the discrimination of most of the newly described species of RGM (38, 101–103).

Partial 16S rRNA Gene Sequence Analysis Generally, the identification of mycobacteria, including RGM, focuses on two main hypervariable domains, known as region A and region B, located at the 5′ end of the 16S rRNA gene. These regions correspond to Escherichia coli positions 129 to 267 and 430 to 500, respectively. Hypervariable region A, especially, contains most of the speciesspecific sequence variations (so-called “signature sequences”) in mycobacterial species, and sequencing of this region allows taxonomic identification of most mycobacteria, including many species of RGM (2, 38). A commercial gene sequencing system, the MicroSeq 500 16S rDNA bacterial sequencing kit (Life Technologies, Foster City, CA), analyzes the first 500-bp sequence and compares the sequence with those in a commercially prepared database. The use of this commercial system alone cannot differentiate some major species of RGM, such as M. chelonae and M. abscessus, which require sequencing of other regions or other genes for identification (101, 103). For this reason, clinical laboratories have supplemented the commercial database with additional sequences from their own or other libraries, such as RIDOM (Würzburg, Germany) or GenBank. The currently commercially available partial 16S rRNA gene sequencing system (MicroSeq ‘first 500 bp’) has a number of large gaps for identification of RGM. For example, isolates of two major RGM pathogens, M. chelonae and M. abscessus, require sequencing of sites outside the first 500 bp, as they are identical in this region but differ at other 16S rRNA gene sites (in the 3′ region) (38, 101, 102). To ensure accurate identification, at least 300 bp of quality sequence should be compared between the reference and query sequences and should cover at least one region of the gene where variations are to be expected. Therefore, most clinical laboratories currently sequence between 450 and 480 bp in order to provide an adequate sequence (104). In general, members of the genus Mycobacterium are closely related to each other, and closely related species may differ by only a few base pairs, or none at all. For example, M. goodii, which is difficult to distinguish from M. smegmatis phenotypically, except by susceptibility pattern, has only a four-base difference in the entire 16S rRNA gene (26). A quality-controlled database is indispensable for the evaluation and accurate identification of unknown strains (103). The laboratorian should also recognize that sequence analysis is an important component in a polyphasic approach to the identification of unknown strains. While in

some instances molecular analysis-based identifications without conventional testing may be adequate, more often there are cases in which the broader picture must be reviewed. Some investigators suggest that key phenotypic tests, including colonial morphology, pigmentation, and growth rate, are necessary, especially for the differentiation of closely related species (103). The lack of consensus on standard reporting criteria or a cutoff value has been a major obstacle in interpretation of sequence data (103, 104). Reporting criteria, such as (i) distinct species, (ii) “related” to a species, and (iii) “most closely related to” a species, depending upon the amount of sequence difference between the unknown isolate and the 16S rRNA gene database entries (40), have been recommended but not validated (38, 103). The 2008 CLSI MM18-A (104) document recommends guidelines for 16S rRNA gene sequencing in order to identify Mycobacterium spp. in a consistent, practical manner. For sequences with 100% sequence probability, the definite genus and species may be assigned. However, for sequence probabilities of 99.0 to 99.9%, the document recommends reporting “genus, most closely related to species,” and for isolates with sequence probabilities of ≥95% to 98.9%, laboratories should consider reporting “unable to definitively identify by 16S rRNA gene sequencing, most closely related to Mycobacterium sp.” (104). We agree that although 100% identity is mandatory for signature sequences, one or a few mismatches at other positions may be acceptable for identification to the species level (104). Sequencing remains a complex and often cost-prohibitive expense for a routine clinical laboratory, which also may not have an adequate volume of isolates to warrant sequencing. Therefore, the consensus opinion is that not all laboratories should attempt to incorporate sequencing into their laboratory routine. Moreover, requests for sequencing should instead be sent to a qualified reference laboratory with skill and experience in the method (103, 104).

Sequencing of the hsp65 Gene Although the 65-kDa heat shock protein gene (hsp65) is highly conserved among species of mycobacteria, it exhibits more interspecies and intraspecies polymorphisms than those in the 16S rRNA gene sequence (105–107). This variability can be advantageous for the development of other strategies for the identification of genetically related species of RGM (106, 107). Most sequencing or restriction fragment length polymorphism analyses have utilized a 441-bp sequence identified by Telenti et al. (107), often referred to as the Telenti fragment. Studies based on DNA sequencing have demonstrated interspecies allelic diversity within the RGM. Detailed studies of several RGM species, including M. peregrinum, M. porcinum, M. senegalense, M. chelonae, and M. abscessus, have shown 4 to 6 sequence variants (sequevars) per species that differ by 4 to 6 nucleotides within the 441-bp Telenti fragment (107, 108). Additionally, unlike 16S rRNA gene sequencing, the hsp65 sequencing method is able to differentiate isolates of M. abscessus from those of M. chelonae (they differ by almost 30 bp in the 441-bp hsp65 sequence, compared to only 4 bp in the entire 1,500-bp 16S rRNA gene sequence) (108). In contrast to the case with 16S rRNA gene sequences, with sequencing of the hsp65 gene, even RGM species with a high degree of 16S rRNA gene similarity, such as M. fortuitum, M. septicum, M. peregrinum, M. houstonense, and M. senegalense, can be discriminated as distinct species.

32. Rapidly Growing Mycobacteria n 603

Like the case for other sequencing methods, one limitation of sequencing of the hsp65 gene is that few or no sequences of newer RGM species are available in databases, and detailed sequencing of older species (i.e., multiple strains) has not been done, such that only one sequence per species is generally available. Thus, development of a comprehensive database and in-house validation are essential (2, 103, 108).

rpoB Sequence Analysis

Initial studies using rpoB (the gene encoding the β-subunit of RNA polymerase) sequencing for description of species were based upon partial rpoB gene sequence analysis, comprising only about 20% of the entire gene length. Other investigators have suggested that species identification of a variety of RGM species is possible using a 340- to 360-bp region, but extensive variation may require development of more species-specific probes (20–22, 109–111). The utility of rpoB was recently emphasized in a study comparing the phylogenetic relationships among 19 RGM species, including the major pathogens in this group, by comparing rpoB to several different sequence targets, including the 16S rRNA gene, hsp65, sodA, and recA. All 19 species showed good discrimination with rpoB (22, 112). Not only has the rpoB sequence been useful for the identification of established species, but it has also helped to enable the discrimination of species that cannot be differentiated by the 16S rRNA gene or the hsp65 sequence alone. Newly described species that are usually differentiated by rpoB sequencing include M. abscessus subsp. massiliense, M. phocaicum, and M. aubagnense (20, 21, 23). However, recent studies have shown that multilocus sequencing is necessary to identify M. abscessus subsp. massiliense (7, 8, 115).

Sequence Analysis of Other Gene Targets Other molecular targets for taxonomic identification, including the 32-kDa protein gene, the superoxide dismutase gene (sodA), the 16S-23S rRNA ITS, dnaJ, secA1, and recA, have been suggested for mycobacterial identification utilizing either PCR-restriction fragment length analysis (PRA) or direct sequencing (22, 51, 113–115). However, preliminary data suggest that these gene sequences are more variable than hsp65, and, to date, they have been utilized less commonly for the laboratory identification of the species of RGM (22, 113, 114). Moreover, a major limitation of all sequence-based testing is the lack of sufficient databases (21, 103). Additionally, a multigenic approach for taxonomic evaluation of species has been suggested widely by investigators and was recently proposed by an ad hoc committee for the reevaluation of the species definition in bacteriology (116). Recent studies by Kim et al. used erm(41) gene sequences to differentiate M. abscessus subsp. massiliense from M. abscessus subsp. abscessus and M. abscessus subsp. bolletii (117). The results of species identification using erm(41) were concordant with those of multilocus sequence analysis including rpoB, hsp65, sodA, and the 16S-23S ITS (117). However, Blauwendraat et al. reported that sequencing of sodA provides little value in species differentiation (98).

Multilocus Sequence Analysis Recent studies have shown the inaccuracy of single-target sequencing for differentiation of species within the M. abscessus complex (i.e., M. abscessus subsp. abscessus, the former species M. massiliense, and the former species M. bolletii)

(118). The previous strategy, using rpoB sequencing for identification of species within the M. abscessus complex, appears to be unacceptable due to the composite genetic structure, which suggests that genetic exchanges among members of the group led to legitimate recombination events among homologous housekeeping genes in a large number of isolates of the M. abscessus complex studied thus far (114, 118, 119). These studies also emphasize the need for multiple-gene sequencing in the definition of species (118). A 2011 study by Macheras et al. suggested the use of eight housekeeping gene sequences (4,071 bp), including argH, cya, glpK, gnd, murC, pgm, pta, and purH (120). Their work also brings into question the previous criteria for differentiation of species based on >3% rpoB sequence divergence between two RGM species (36, 120). Furthermore, their work substantiates a later study by Leao and colleagues asserting that M. massiliense and M. bolletii are not different species (7, 8).

PCR-Restriction Enzyme Analysis PRA of hsp65 has become a valuable tool for the identification of RGM. With the nonsequencing PRA method used on hsp65, minor differences (sequevars) within the species rarely involve a restriction site, so most species have only one PRA pattern. Currently, the 441-bp Telenti fragment of hsp65 remains the most useful sequence for PRA identification of RGM, although it has not been evaluated extensively with the pigmented RGM and with the newer species and subspecies of RGM, such as M. phocaicum, M. aubagnense, M. abscessus subsp. massiliense, and others (1, 107, 108). However, use of another restriction enzyme (SmlI) in addition to the BstEII and HaeIII enzymes may enable differentiation of M. abscessus subsp. massiliense from the former species M. bolletii and M. abscessus (R. J. Wallace, unpublished data). Additionally, the correlation of rough/ smooth colony morphology of M. abscessus subsp. massiliense has been demonstrated by hsp65 PRA with Hinf1 (13). The advantages of PRA are that the method of identification does not rely upon growth rate and nutritional requirements, the equipment is relatively inexpensive, and the results for a large number of mycobacterial species can be generated rapidly. The disadvantages are that it requires knowledge of PCR and is a relatively complex procedure that requires extensive in-house validation, since there are no available commercial systems. Furthermore, the method is not approved by the U.S. Food and Drug Administration. However, as with all sequence-based methods of identification, its utility is limited by the availability of an updated public database. Algorithms for identification of mycobacterial species, including RGM, by PCR-restriction fragment length polymorphism analysis of the hsp65 gene have been proposed (89, 106, 107). Figure 1 shows a PRA gel of the most commonly encountered RGM in the clinical laboratory.

Variable-Number-Tandem-Repeat (VNTR) Analysis Recently, Choi and colleagues demonstrated identification to the species level of 85 isolates of the M. abscessus complex (M. abscessus subsp. abscessus, M. abscessus subsp. massiliense, and M. abscessus subsp. bolletii), using VNTR11 and VNTR23 as targets (121). The assay was also able to differentiate the M. abscessus complex from other mycobacterial species, including M. fortuitum and M. chelonae. Complete agreement of identification was noted among the 85 strains. VNTR11 showed a polymorphism of two to four copies (179 to 278 bp) in M. abscessus but only one copy in

604 n

BACTERIOLOGY

The sequence, located in variable region II according to the numbering of Adékambi et al. (20), at positions 1105 to 1219 (including primer sequences), successfully identified 99/100 isolates of M. abscessus and M. chelonae (125).

Matrix-Assisted Laser Desorption Ionization—Time of Flight Mass Spectrometry

FIGURE 1 PRA patterns of commonly encountered species of RGM (106, 107). Lane 1, M. chelonae, BstEII; lane 2, M. abscessus subsp. massiliense, BstEII; lane 3, M. abscessus subsp. abscessus, BstEII; lane 4, M. fortuitum, BstEII; lane 5, 100-bp ladder; lane 6, pGem ladder; lane 7, M. chelonae, HaeIII; lane 8, M. abscessus subsp. massiliense, HaeIII; lane 9, M. abscessus subsp. abscessus, HaeIII; lane 10, M. fortuitum, HaeIII. doi:10.1128/9781555817381.ch32.f1

M. abscessus subsp. massiliense and M. abscessus subsp. bolletii. VNTR23 was present as one to three copies (196 to 238 bp) in M. abscessus, in contrast to one pattern with two copies in M. abscessus subsp. bolletii. None of the M. abscessus subsp. massiliense isolates were positive for VNTR23 (121).

Pyrosequencing Pyrosequencing technology (Biotage, Uppsala, Sweden) employs nucleic acid sequencing of a 20- to 30-bp segment of hypervariable region A of the 16S rRNA gene. The method is based on the detection of pyrophosphate during DNA synthesis. During sequencing, visible light is produced that is proportional to the number of incorporated nucleotides. The method is not as discriminating as traditional Sanger sequencing but is an attractive and less expensive alternative. Another advantage of the pyrosequencing technology is its commercial availability. Like the case for other sequencing methods, the major limitation is the quality of the databases for interpretation and comparison of sequences (122). In a recent study of 50 RGM isolates (M. chelonae/ M. abscessus and M. mucogenicum), consensus sequences were obtained for 40 isolates compared to traditional sequencing results. Of 10 isolates of M. fortuitum, 3 had identical sequences which matched those of M. fortuitum/M. peregrinum, and the remaining 7 isolates matched M. fortuitum (123). To date, this method has provided reliable and rapid identification of a variety of RGM species (123). A real-time PCR with melting curve analysis has been developed that consistently detects and differentiates M. tuberculosis from NTM. In a recent study, 20 isolates previously identified as M. fortuitum were confirmed to be M. fortuitum. However, 7 of 24 isolates previously identified as M. chelonae or M. abscessus were found by pyrosequencing analysis to have been misidentified by traditional methods (124). The application of these methods is currently used only in research or large reference laboratories, until more extensive evaluation can be done. Recently, investigators proposed the use of a segment of the rpoB gene in conjunction with pyrosequencing to determine the sequence of a 60-bp portion of the rpoB gene.

Matrix-assisted laser desorption ionization—time of flight mass spectrometry (MALDI-TOF MS) technology was recently demonstrated to be a reliable and rapid (approximately 90 min to identification of most RGM species) technological advance for the identification of NTM, including the RGM. The current method produces unique spectral fingerprints based on extracted proteins. Moreover, MALDI-TOF MS provides a less expensive and less labor-intensive procedural alternative to gene sequencing for identification of the RGM (126, 127). In a recent study by Lotz and colleagues, the investigators were able to differentiate several clinically relevant species of RGM, including M. immunogenum/M. chelonae/M. abscessus subsp. abscessus, but not M. abscessus subsp. abscessus, from M. abscessus subsp. bolletii or M. abscessus subsp. massiliense (128). Additionally, 6/8 isolates of M. fortuitum, 8/8 isolates of M. mucogenicum, and 6/7 isolates of M. peregrinum were also correctly identified in liquid media, compared to 25/ 26 isolates of M. chelonae, 25/26 isolates of M. fortuitum, 13/16 isolates of M. peregrinum, and 19/20 isolates of M. mucogenicum correctly identified from solid media (128). Overall identification for both slowly and rapidly growing mycobacterial species was 97% (128). The original studies using MALDI-TOF MS began in 2004, using more complex strategies, with whole cells for mycobacterial identification, than the protein extract methods available at present. In early studies, Hettick et al. tested eight strains of RGM (M. fortuitum) and were able to illustrate a high degree of reproducibility between culture strains (129, 130). Additionally, Wallace and colleagues tested 22 isolates of RGM and showed that the clustering of strains by MALDI-TOF MS was comparable to that by PRA and genetic sequencing (18). In 2006, Pignone et al. used root mean square values for the comparison of different mycobacterial profiles of 14 strains of RGM (5 M. fortuitum group, 3 M. mucogenicum, 3 M. abscessus, and 3 M. chelonae strains) (131). These investigators showed reproducible identification, with unique mass spectral profiles of all 14 strains of the RGM, extending the work of the previous studies by Hettick et al. (129, 131). Although Lefmann et al. identified 24 clinical and 12 type strains of mycobacteria (including 9 strains of the M. tuberculosis complex) by using MALDI-TOF MS, they only tested two clinical and two type strains of RGM, and their strategy entailed initial PCR and transcription-RNase cleavage steps, unlike the systems currently being evaluated (132). Today’s MALDI-TOF MS systems are more easily incorporated into the routine workflow of the laboratory and provide rapid and accurate identification of many strains of mycobacteria, as shown by Saleeb and colleagues (126). Currently, the two predominant commercial MALDI-TOF MS systems are the Bruker Biotyper (Bruker Daltonics, Billerica, MA) and bioMerieux Vitek MS (bioMerieux, Durham, NC) systems. The current systems also contain a spectral database and identification algorithms for the detection of conserved and microbe-specific peak patterns in wholecell mass spectra (127). The most time-consuming portion of the current method for identification of mycobacteria is the inactivation and extraction procedure (126). Although MALDI-TOF MS is able to identify multiple species of

32. Rapidly Growing Mycobacteria n

RGM, the method is still inadequate to distinguish closely related RGM species, including M. abscessus subsp. abscessus versus M. abscessus subsp. massiliense, and M. mucogenicum versus M. phocaicum (126). Just as for genetic sequencing methods, the importance of an extensive database for MALDI-TOF MS cannot be overemphasized. Most users are currently supplementing the available commercial databases with their own in-housedeveloped databases to allow species-level differentiation of a greater number of species of NTM (126). Future applications suggested for MALDI-TOF MS include refinement of the databases, possible typing of organisms, detection of the absence or presence of specific proteins associated with virulence and antimicrobial resistance, and development of methods for direct detection and identification from clinical samples (127).

TYPING SYSTEMS Pulsed-Field Gel Electrophoresis Pulsed-field gel electrophoresis (PFGE) is the most widely used method for molecular strain typing of the RGM. Although PFGE has never been standardized for RGM, most investigators concur that small (2 or 3 bands) differences between isolates indicate that the isolates are closely related; differences of 4 to 6 bands indicate that the strains are possibly related, and differences of ≥7 bands indicate that the isolates are genetically different (16, 82, 84, 133). Because unrelated strains of most RGM contain highly diverse PFGE patterns, this technique has been useful in epidemiological investigations. With the addition of thiourea as a modification of the original method, it is now possible to obtain reliable results by PFGE for all species of RGM, including isolates previously affected by DNA degradation (134, 135). Recent studies have shown that DNA smear patterns of previous isolates are correlated with the presence of dnd, a DNA degradation gene (136).

Random Amplified Polymorphic DNA PCR In the random amplified polymorphic DNA PCR (RAPDPCR) method, using one arbitrary primer and low-stringency conditions, the primer hybridizes to both strands of template DNA where it is matched or partially matched, resulting in strain-specific heterogeneous DNA products. Zhang and colleagues applied the RAPD-PCR or arbitrarily primed PCR analysis method to compare strains of M. abscessus (134). They were able to confirm several previous observations about prior nosocomial RGM outbreaks, including a 1988 epidemic of otitis media due to aminoglycoside-resistant M. abscessus in children with prior tympanostomy tubes, as well as a cardiac surgery outbreak (1, 134).

VNTR Analysis The development of strain typing methods using VNTR would be valuable for helping to explain differences between isolates from chronic new infections and relapse infections. Blauwendraat and colleagues also indicated that the method would show whether or not cross-infection occurs and, moreover, enable prospective studies of strains of the M. abscessus complex correlated with clinical outcomes (98). Recently, Wong et al. (137) described a VNTR typing assay (MavA) using 18 tandem repeats in the M. abscessus genomic sequence, which was found to be superior to multilocus sequencing, with 100% typeability, 100% locus stability, and 100% reproducibility.

605

Repetitive-Sequence-Based PCR A commercial system, DiversiLab system (bioMerieux, Durham, NC), is available for strain typing of organisms, including mycobacteria, by using repetitive elements interspersed throughout the genome. The system electrophoretically separates repetitive-sequence-based PCR (rep-PCR) amplicons on microfluidic chips to provide computer-generated readouts. The discriminative power has been reported to equal or exceed that of standard restriction fragment length polymorphism analysis for some species of mycobacteria, with a smaller sample size than that for standard PFGE, and in a much shorter time frame (114, 138). Limitations of the system include the lack of an extensive established database and the cost of the system.

Enterobacterial Repetitive Intergenic Consensus PCR (ERIC PCR) Enterobacterial repetitive consensus sequences are repetitive elements distributed along the bacterial chromosome, at intergenic regions of polycistronic operons or flanking open reading frames. The method was recently evaluated with isolates of the M. abscessus/M. chelonae complex and with isolates of M. fortuitum (87, 139). Typing of isolates by ERIC PCR works in mycobacteria as a RAPD PCR because the presence of ERIC repeats has never been demonstrated in available Mycobacterium genomes, and amplification with appropriate ERIC primers can occur in the absence of genuine ERIC sequences (139). In a study of outbreak strains of M. abscessus in Brazil, ERIC PCR showed higher discriminatory power than PFGE for typing of strains which had shown smear patterns with PFGE using thiourea (139), although this method was not as discriminatory when testing isolates of M. fortuitum (87, 139).

SEROLOGIC TESTS Serologic classification of mycobacteria has been attempted starting in 1925 with M. avium complex. However, serotyping has not been suitable for routine species identification of mycobacteria including the RGM and early studies served to emphasize the complexity of the antigenic composition of mycobacteria with many antigens shared by more than one species (140).

ANTIMICROBIAL SUSCEPTIBILITIES Several different methods have been used for susceptibility testing of RGM for clinical purposes. These methods include agar disk diffusion, broth microdilution, agar disk elution and E-test. Each method has proved useful but none of the methods were well standardized until 2003 with the publication of the Clinical and Laboratory Standards Institute (CLSI) guidelines (141). In the M24A document, the CLSI recommended broth microdilution as the reference method for susceptibility testing of the RGM (142). Nine antimicrobials including amikacin, cefoxitin, ciprofloxacin, clarithromycin, doxycycline, linezolid, imipenem, sulfamethoxazole and tobramycin have been recommended for testing and breakpoints have been established for these agents. A recent tentative CLSI guideline revision to include additional agents such as moxifloxacin, meropenem, minocycline, and trimethoprim-sulfamethoxazole has also been published (142). Additionally, a change of the imipenem intermediate breakpoint from 8 μg/ml to 8–16 μg/ml has enabled reporting of this drug with isolates of M. abscessus and M. chelonae which was not previously recommended (142).

606 n

BACTERIOLOGY

Briefly for the broth microdilution method, drug dilutions are prepared using serial 2-fold dilutions of cationadjusted Mueller-Hinton broth. Suspensions of organisms are prepared to match a 0.5 McFarland turbidity standard. The suspensions are then diluted to a concentration of approximately 106 CFU/ml. From that suspension, 100 μl is delivered into the wells of a 16-well microtiter plate with a final concentration of approximately 10 4 CFU/well (142). MICs are optimally read after incubation at 30°C for 3 days. Several specific recommendations about test results have also been made. Tobramycin should only be reported for isolates of M. chelonae. Any RGM isolate with an amikacin MIC of ≥64 μg/ml should be retested and/or sent to a reference laboratory in order to confirm resistance (although mutational resistance involving the 16S rRNA gene does occur). Also, if any isolate of the M. fortuitum group has an imipenem MIC >8 μg/ml, this should also be repeated with careful attention paid to inoculum density and a maximum incubation time of 3 days because of the instability of imipenem over time. Another caveat is that the MICs of sulfamethoxazole and trimethoprim sulfamethoxazole are read using 80% inhibition of growth as the susceptibility endpoint not 100% inhibition as is used for the other antimicrobials. Overinoculation of the MIC panels is often most obvious with sulfonamides. An inexperienced laboratorian may interpret the sulfonamide MIC as resistant when in reality the inoculum was too heavy. Rarely isolates of the M. fortuitum group are resistant to sulfonamides. If an isolate in this group is found resistant, a repeat test with lower inoculum is warranted (142). The recent finding of the presence of a functional erm gene that induces macrolide resistance in many isolates of the M. fortuitum group, and M. abscessus subsp. abscessus but not M. chelonae or M. abscessus subsp. massiliense also has made changes to the manner in which clarithromycin reporting should be done (94, 95). A major revision to the CLSI document has been added to recommend an initial 3-day reading of MICs followed by a final reading at 14 days (unless the isolate becomes resistant before that time) to detect a functional erm gene in RGM isolates (142). The significance of the finding of these genes has not yet been assessed in clinical trials. However, a recent landmark study in Korea among 64 patients with M. abscessus subsp. abscessus and 81 patients with M. abscessus subsp. massiliense lung disease showed that approximately 80% of the patients with M. abscessus subsp. massiliense could be successfully treated with a macrolide containing multidrug regimen in contrast to only about 20% of the patients with M. abscessus subsp. abscessus (64). The investigators suggested that inducible resistance to clarithromycin could explain the decreased efficacy of clarithromycin containing antibiotic regimens against M. abscessus lung disease (64). A 2012 study by Choi and colleagues has also shown other susceptibility differences between M. abscessus subsp. massiliense (143). These investigators observed evidence of in vitro, ex vivo, and in vivo activity of moxifloxacin and two macrolides, clarithromycin and azithromycin against M. abscessus and M. abscessus subsp. massiliense (143). When moxifloxacin was combined with a macrolide against M. abscessus, antagonism (defined as fractional inhibitory concentration (FIC) ≥2) was observed in 65.4% (7/26) strains with clarithromycin and 46.2% (12/26) with azithromycin in vitro. Similar results were seen in macrophage cultures and a murine model. In contrast, however, either indifferent (FIC >0.5 and 80 named species distributed in 17 genera, of which 4 are known to contain human pathogens: Actinobacillus, Aggregatibacter,

Capnocytophaga The Capnocytophaga genus in the family Flavobacteriaceae at present consists of nine species (C. canimorsus, C. cynodegmi, C. ochracea, C. gingivalis, C. sputigena, C. haemolytica, C. granulosa, C. leadbetteri, and genospecies AHN8471) of

*This chapter contains information presented by Reinhard Zbinden and Alexander von Graevenitz in chapter 33 of the 10th edition of this Manual.

doi:10.1128/9781555817381.ch35

652

35. Fastidious Gram-Negative Rods n 653

facultatively anaerobic, nonmotile, Gram-negative rods, but with gliding motility (14). The G+C content of the DNA of this genus is 34 to 44 mol% (15).

S. moniliformis, with a G+C content in its DNA of 24 to 26% mol% (27). See chapter 54 for the genus Leptotrichia of the family Leptotrichiaceae.

Cardiobacterium

Suttonella

The Cardiobacterium genus in the family Cardiobacteriaceae consists of facultatively anaerobic, nonmotile, Gramnegative rods, with the species C. hominis and C. valvarum (16). The G+C content of the DNA of these species is 59 to 60 mol% (17).

The Suttonella genus in the family Cardiobacteriaceae (17) consists of facultatively anaerobic, nonmotile, Gramnegative rods and so far contains only one species, S. indologenes (formerly Kingella indologenes) (http://www.bacterio .net). The G+C content of its DNA is 49 mol% (17).

Chromobacterium The Chromobacterium genus in the family Neisseriaceae (18) contains several facultatively anaerobic, motile species, of which C. violaceum is at present the only agent of human disease. Its DNA content is between 65 and 68 mol% (18). C. haemolyticum, isolated from a sputum culture, is so far represented by one strain only (19) and is not covered here.

Dysgonomonas The genus Dysgonomonas in the family Porphyromonadaceae consists of facultatively anaerobic, nonmotile, Gramnegative rods (20). Four species have been described: D. capnocytophagoides and D. gadei (21), D. mossii (22), and D. hofstadii (23). The G+C content of the DNA is ∼38 mol% (20).

Eikenella The genus Eikenella in the family Neisseriaceae (18) consists of facultatively anaerobic, nonmotile, Gram-negative rods. Thus far, only one species, E. corrodens, has been recognized. The G+C content of the DNA of Eikenella is between 56 and 58 mol% (18).

Kingella The genus Kingella in the family Neisseriaceae consists of the facultatively anaerobic, nonmotile species K. kingae, K. denitrificans, K. oralis, and K. potus (18, 24). The G+C content of the DNA of these species is between 47 and 58 mol% (18).

Pasteurella The taxonomy of the genus Pasteurella in the family Pasteurellaceae has been in flux for some time (10, 13; http:// www.bacterio.net). P. multocida can be separated into the subspecies multocida, septica, and gallicida; P. canis, P. dagmatis, and P. stomatis are other species isolated from humans. In spite of the genotypical homogeneity of P. multocida isolates, phenotypically diverse lineages have been observed, e.g., sucrose-negative variants from infections from bite wounds made by large cats (25). The G+C content of the DNA of Pasteurella species is between 38 and 46 mol% (13). New genera or reclassifications may be necessary for species discussed below that are preceded by “(P.)” or “(Pasteurella)” (26).

Simonsiella The genus Simonsiella in the family Neisseriaceae consists of several obligately aerobic species that may show gliding motility (18). The only species isolated from humans is S. muelleri. This genus has a G+C content in its DNA of 40 to 50 mol% (18).

Streptobacillus The Streptobacillus genus in the family Leptotrichiaceae (27) consists of one facultatively anaerobic, nonmotile species,

EPIDEMIOLOGY AND TRANSMISSION Most bacteria in this group are part of the microbiota of the nasopharynx and/or the oral cavity of animals and/or humans and are parasitic, with the only environmental genus being Chromobacterium. Transmission from animals occurs by contact (e.g., bites and licking of wounds), from humans to humans by droplets (e.g., directly with Kingella spp. or by paraphernalia or human bites with E. corrodens). They may cause infections anywhere in the human body. Risk factors exist for certain types of septicemia (e.g., liver cirrhosis for P. multocida, neutropenia for oxidase-negative Capnocytophaga spp., and chronic granulomatous disease for C. violaceum). Endogenous infections occur as well, e.g., HACEK (Haemophilus spp., Aggregatibacter spp., Cardiobacterium spp., E. corrodens, and Kingella spp.) endocarditis (28). The habitat of the HACEK group, e.g., Haemophilus spp. and Aggregatibacter spp., is the human oral cavity, including dental plaque; infections are endogenous (11, 29). The normal habitat of Cardiobacterium spp. is the human oral cavity and nasopharynx but possibly also the gastrointestinal and urogenital tracts (17, 28). Infections are endogenous. The natural habitat of E. corrodens is the oral cavities and possibly the gastrointestinal tracts of humans and some mammals, from which it can be transmitted via saliva (bites, syringes) to other individuals (30–32). Endogenous infections prevail, however. The natural habitat of Kingella spp. is the upper respiratory tract and oral mucosa of humans and possibly other primates. K. kingae colonizes the throat but not the nasopharynx of many children aged 6 months to 4 years (33). Ribotyping and pulsed-field gel electrophoresis have shown that K. kingae can be transmitted via respiratory droplets (34), although most infections are endogenous. K. oralis has been isolated from the human mouth (18). S. indologenes is not an official constituent of HACEK but is well known as an agent of endocarditis (35). However, its natural habitat is not known (17). A. lignieresii (primary habitat in the oral cavities of sheep and cattle), A. equuli (in the oral cavities of horses and pigs), and A. suis (in the oral cavities of pigs) can be transmitted to humans by animal contact (29). Exclusively human are A. hominis and A. ureae, whose normal habitat is unknown (29, 36). The oxidase- and catalase-negative species C. ochracea, C. gingivalis, C. sputigena, C. haemolytica, and C. granulosa, as well as the recently described C. leadbetteri and genospecies AHN8471, are normal but not prominent members of the human oral microbiota (14). The first three have been isolated from adults with periodontal disease but also from periodontitis-free adults; the other four have been isolated from supragingival and subgingival plaque in children and adults (14). Infections are endogenous. The oxidase- and catalase-positive species C. canimorsus and C. cynodegmi reside in the oral cavities of healthy dogs (25% of dogs have C. canimorsus as determined by culture and 85 to 100% of

654 n

BACTERIOLOGY

dogs have it as determined by PCR) and cats (15%, as determined by culture) (37). Chromobacterium inhabits soil and water in tropical and subtropical climates between latitudes of 35°N and 35°S (South Africa, Southeast Asia, Australia, southeastern United States, and, rarely, South America) (38, 39). The portal of entry is usually the skin, but oral intake has also been reported (40). Most D. capnocytophagoides strains have been isolated from stools of immunocompromised patients, and a few strains have been isolated from other sources (41). The natural habitats of this and the other Dysgonomonas spp. are unknown (22, 23). Pasteurella spp. are widespread in healthy and diseased wild and domestic animals, including rodents, dogs, and cats, inhabiting the nasopharynx and gingiva (13). Human isolates are transmitted predominantly from animals by contact (bites or licking or scratching of wounds). Of the “related” species, (Pasteurella) aerogenes occurs primarily in pigs (42), (Pasteurella) caballi in pigs and equines (42, 43), and (Pasteurella) pneumotropica in rodents and dogs (44); the natural habitat of (Pasteurella) bettyae is uncertain. The natural habitat of S. muelleri is the oral cavity of humans. In healthy human populations, the incidence of Simonsiella spp. is in the range of 30 to 40%. Children possibly have a higher incidence than adults. In dogs and cats, Simonsiella spp. are common and abundant (18). S. moniliformis occurs naturally in the upper respiratory tract of up to 100% of wild and laboratory rats and other rodents (mice, gerbils, squirrels, ferrets, and weasels) and occasionally of dogs and cats preying on rodents. Transmission to humans occurs either from bites of those animals (rat bite fever) or from consumption of contaminated food or water (Haverhill fever) (45).

CLINICAL SIGNIFICANCE Actinobacillus spp. A. lignieresii causes actinobacillosis, a granulomatous disease in cattle and sheep in which, similar to actinomycosis, sulfur granules form in tissues (46). A few human soft tissue infections after a cow or sheep bite or other contacts have been reported (42). A. equuli and A. suis have caused a variety of diseases in horses and pigs; human infections are generally due to horse or pig bites or contact (42). Both species have also been isolated, albeit rarely, from the human upper respiratory tract (13, 47). A. ureae is most often a commensal in the human respiratory tract, particularly in patients with lower respiratory tract disease (13), but has also been found as an agent of meningitis following trauma or surgery (48) and of other infections in immunocompromised patients (49). A. hominis has also been isolated from such patients but has occurred as a commensal as well, albeit rarely (36). Virulence factors belong to the pore-forming protein toxins of the RTX family; RTX toxins have repeats in the structural toxin peptide and exhibit a cytotoxic and often also a hemolytic activity. They are particularly widespread in species of the family Pasteurellaceae (50).

Aggregatibacter spp. A. actinomycetemcomitans is one of the major agents of juvenile and adult periodontitis (51) and may occur together with Actinomyces spp. in actinomycotic sulfur granules (11). Furthermore, it may cause HACEK endocarditis (28), soft tissue infections, and other infections (52). In a large, multinational cohort study, HACEK organisms were the causes

of 1.4% of all cases of endocarditis (53). HACEK endocarditis is characterized by a relatively long interval between first symptoms and diagnosis (range, 2 weeks to 6 months), large vegetations on native or artificial valves of the left side, and frequent embolizations. Prognosis is good with appropriate antibiotic treatment (28, 53). Virulence factors are an RTX leukotoxin (50), a cytotoxic distending toxin (54), and the adhesin EmaA (55), as well as fimbriae (56). A. aphrophilus may cause systemic disease, particularly bone and joint infections, spondylodiscitis, and endocarditis (52, 57, 58). A. segnis, whose frequency may be underestimated due to apparent misdiagnoses, may cause endocarditis (59). It may be an important cause of bacteremia and sometimes of other infections, e.g., pyelonephritis (60).

Capnocytophaga spp. C. ochracea, C. gingivalis, C. sputigena, C. haemolytica, and C. granulosa have been reported as agents of septicemia and other endogenous infections (endocarditis, endometritis, osteomyelitis, soft tissue infections, peritonitis, ophthalmic lesions, and noma) (30, 47, 61–63) in immunocompetent and immunosuppressed (mainly neutropenic) patients. They are able to suppress neutrophilic chemotaxis and lymphocyte proliferation (64). The association with periodontitis remains unclear (14). Infections with C. canimorsus and C. cynodegmi are associated mainly with dog or cat bites or contact. Patients infected with C. canimorsus most often present with septicemia and have previously been splenectomized or are alcoholics. In fulminant cases with a poor prognosis, disseminated intravascular coagulation, acute renal failure, respiratory distress syndrome, and shock may develop (65). Hemolyticuremic syndrome and thrombotic thrombocytopenic purpura are other possible sequelae (66). Meningitis (67, 68), keratitis (65), and endocarditis (69) have been reported as well. C. cynodegmi has been isolated more rarely, mainly from localized or systemic infections (70). C. canimorsus resists phagocytosis by macrophages and killing by complement and leukocytes; macrophages incubated with the bacterium fail to produce several proinflammatory cytokines (71).

Cardiobacterium spp. Disease caused by both Cardiobacterium species is mainly HACEK endocarditis (16, 53); C. hominis prosthetic valve endocarditis has been described after upper endoscopy and colonoscopy (72), and on rare occasions, C. hominis has been isolated from other body sites (30, 73). Most reported cases of C. valvarum endocarditis are related to periodontal diseases (74). In blood culture-negative cases, the diagnosis has been made by broad-range PCR applied to valve tissue (75).

Chromobacterium violaceum Localized infections usually arise from contaminated wounds, and septicemia with multiple organ abscesses may follow. They are significantly associated with neutrophil dysfunction (glucose-6-phosphate dehydrogenase deficiency, chronic granulomatous disease). Children without these conditions and those with bacteremia show a high fatality rate (38–40). A number of virulence factors other than endotoxin, i.e., adhesins, invasins, and cytolytic proteins, have been described (76).

Dysgonomonas spp. Diarrhea was reported to have occurred in 10 of 20 patients with fecal isolates of C. capnocytophagoides, whereas routine

35. Fastidious Gram-Negative Rods n 655

stool cultures yielded the organism in 1.1 to 2.3% of cultures (77). Bacteremia occurs as well (41); one blood isolate was found to be identical by ribotyping to one in the stool of the same patient (78). One strain of D. mossii was isolated from intestinal juice of a patient with pancreatic carcinoma (79). D. gadei has been isolated from a human gallbladder (21) and from blood (80), and D. hofstadii has been isolated from a wound (23).

Eikenella corrodens E. corrodens is associated with juvenile and adult periodontitis (51) but is also an agent of infections of the upper respiratory tract, pleura and lungs, abdomen, joints, bones, wounds (e.g., from a human bite), and, rarely, other infections, like noma (30–32, 81). These organisms are often indolent and found mixed with other members of the oropharyngeal microbiota, particularly staphylococci and streptococci. Risk factors are dental manipulations and intravenous drug abuse. Endocarditis is of the HACEK type if monomicrobial, but polymicrobial non-HACEK cases are known (28). E. corrodens can trigger a cascade of events that induce inflammation in periodontal tissue (82).

Kingella spp. Infections with K. kingae show a predilection for bones and joints of previously healthy children 90% of strains. d Delayed development of hemolysis occurs in 11 to 89% of strains. e Elevated concentrations of CO2 of ≥10% enhance growth. f Beta-hemolysis is produced on horse and sheep blood agar; occasional strains fail to produce beta-hemolysis.

vb − − vb − − uk − − vb + + vb − + uk + + vb − v vb − − uk − + H. H. H. H. H. H. H. H. H.

influenzae aegyptius haemolyticus parainfluenzae ducreyi parahaemolyticus pittmaniae paraphrohaemolyticuse sputorum

+ + + − + − − − −

+ + + + − + + + +

+ + + v − v w + uk

+ +c + + v + + + +

− − − + − + + + uk

− − − − − − − − −

+ − v − − − − − −

− − − + − − + − −

Urease Xylose X factor V factor Catalase Glucose

Sucrose

Lactose

Fermentation of: Requirement for: Haemophilus species

Differential characteristics of Haemophilus speciesa TABLE 1

strain type, the capsule is composed of ribose and fructose in the furanose ring and Glc, Gal, GlcNAc, or ManANAc in the pyranose ring. The structures of the capsules belonging to each serotype can be found in reference 5. Indole, ornithine decarboxylase, and urease production are the basis for a biotyping scheme with both H. influenzae and H. parainfluenzae (11) (Table 2).

EPIDEMIOLOGY AND TRANSMISSION

Indole

Ornithine decarboxylase

β-Galactosidase

BACTERIOLOGY

Mannose

Production of:

668 n

Haemophilus species can be found as part of the commensal biota of mucous membranes in humans. Colonization of the oral cavity superior to the palatal arches by H. parainfluenzae and H. pittmaniae is normal (5, 12–15). H. parahaemolyticus and H. haemolyticus colonization in healthy individuals remains rare, although H. haemolyticus has been isolated from subgingival dental plaques (5). Colonization of the cervix with H. ducreyi has been documented following sexual intercourse (16, 17). H. influenzae may also be found as part of the commensal bacterial biota of the mucosal surfaces of the upper respiratory tract (URT) of many healthy individuals (18); however, asymptomatic colonization of the URT with encapsulated strains of H. influenzae type b (Hib) is rare. Carriage of Hib was reported in only 2 to 5% of healthy children in the prevaccine era and has further decreased (∼0.06%) since the introduction of the pediatric Hib conjugate antigen vaccine, HIB, in the 1980s (19). In contrast, NTHi, together with strains of H. parainfluenzae, represents a major portion of the cultured bacterial microbiota of the pharynx and nasopharynx of >90% of healthy individuals (13, 20). Clones of NTHi present in the URT differ when asymptomatic carriers are compared to those with infection (20, 21). In asymptomatically colonized individuals, the clones vary continuously, with a mean duration of carriage of 1 to 2 months (21). However, during infection, a single clonal group predominates. As a result of widespread use of HIB, the distribution of serotypes causing invasive infection has shifted. A recent report by the U.S. Centers for Disease Control and Prevention (CDC) classified 69.5% of invasive isolates as nontypeable, 2.2% as type a, 3.6% as type b, 0.3% as type c, 0.3% as type d, 5.7% as type e, and 18.3% as type f from 1999 to 2008 (22). Despite the reduction in Hib carriage and the efficacy of the vaccine in preventing invasive infections, the prevalence of invasive H. influenzae and Hib remains high among Alaskan Natives and Native Americans (22). Recently, an increase in the incidence of invasive H. influenzae infections has been observed in patients aged >65 years (22, 23). MacNeil et al. (22) reported that from 1999 to 2008 the incidence of invasive H. influenzae was 4.09 per 100,000 population, with 19.5% of invasive infections occurring in nursing home residents (22). The incubation period for H. influenzae is poorly understood. The presence of a concomitant or preceding viral infection can predispose previously healthy carriers to infection. In these instances, the colonizing bacteria invade the inflamed or damaged mucosa and enter the bloodstream. The antiphagocytic nature of the Hib capsule and the absence of the anticapsular antibody lead to increasing bacterial proliferation (18). When the bacterial concentration exceeds a critical level, it can disseminate to various sites, including the meninges, subcutaneous tissue, joints, pleura, pericardia, and lungs. The presence of antibody, complement, and phagocytic cells determines the clearance of the bacteremia and can influence dissemination (18). Host defenses include activation of the alternative and classical complement pathways and production of antibodies

36. Haemophilus

n

669

FIGURE 1 Electron micrographs depicting an encapsulated type b strain (left) and a nonencapsulated, nontypeable strain (right) of Haemophilus influenzae. doi:10.1128/9781555817381.ch36.f1

directed against the polyribosylribitol phosphate (PRP) capsule of Hib. Antibody reactive with the Hib capsule plays a primary role in conferring immunity. Newborns have a low risk of infection because of the presence of maternal antibodies acquired through colostrum. Beginning at about 2 months of age, when maternal antibodies to the PRP capsule begin to wane, infants are at high risk for developing invasive H. influenzae disease. In the absence of maternal antibodies, and even following natural infection, the antiPRP immune response of infants is diminished (18). Therefore, infants are at risk for repeat infections because prior episodes of infection do not confer immunity. By the age of 5 years, most children have naturally acquired antibodies.

TABLE 2 Biotypes of Haemophilus influenzae and Haemophilus parainfluenzae Production of: Species

H. influenzae

H. parainfluenzae

Biotype

I II III IV V VI VII VIII I II III IV V VI VII VIII

Indole

Urease

Ornithine decarboxylase

+ + − − + − + − − − − + − + + +

+ + + + − − − − − + + + − − + −

+ − − + + + − − + + − + − + − −

CLINICAL SIGNIFICANCE H. influenzae Invasive infections caused by H. influenzae, such as meningitis, epiglottitis, orbital cellulitis, and bacteremia, are usually caused by capsular type b strains and generally fall within biotypes I and II of this species (24). Life-threatening Haemophilus infections, however, have fortunately become exceedingly uncommon in developed countries since the development and introduction of the pediatric HIB vaccine (25–27). When infections caused by Hib occur today, it is usually in the setting of an unvaccinated child, although they may also arise in both children and adults as a result of head trauma or cerebrospinal fluid (CSF) leak or following a neurosurgical procedure. Biotype IV strains, at least in the pre-HIB vaccine era, were often found to cause systemic infections in neonates as well as aggressive infections of the genital tract in postpartum women (28, 29). The vast majority of H. influenzae infections today are caused by NTHi (10, 20). This organism is an important cause of acute conjunctivitis, acute otitis media, acute maxillary sinusitis, acute bacterial exacerbation of chronic bronchitis, and pneumonia (18). The organism gains access to the site of infection by direct contiguous spread from its reservoir in the URT. Spread via respiratory secretions, usually on the hands of patients, NTHi can also lead to conjunctival infection. Antecedent viral infections with resultant inflammation of the eustachian tubes and sinus ostea predispose patients to infection of the middle ear cavity and maxillary sinuses, respectively, by compromising egress from and ingress to these closed spaces (30). Establishment of infection in the lungs is facilitated by any condition that diminishes mucociliary clearance of organisms from the respiratory tree (18, 31, 32). Examples include smoking, chronic obstructive pulmonary disease, viral infection, recurrent bacterial infection, and physiological alterations, such as those that occur in individuals with cystic fibrosis

670 n BACTERIOLOGY

(18, 31, 32). Persons at risk for systemic NTHi infection, particularly the elderly, also include those with functional or anatomic asplenism, sickle cell disease, complement deficiencies, Hodgkin’s lymphoma, congenital or acquired hypogammaglobulinemia, and T-cell immunodeficiency states (e.g., HIV infection). Rarely is NTHi documented to be a cause of bacteremia; in a recent study, Laupland et al. (33) found the incidence of NTHi bacteremia to be 304 cases per 33,601,000 person-years. This may be due to the relative avirulence of the organism, the inadequacy of conventional blood culture techniques in propagating this fastidious bacterium, or the lack of contemporary studies investigating the epidemiology of invasive H. influenzae infections.

H. ducreyi Chancroid is a sexually transmitted disease caused by H. ducreyi that is usually characterized by the development of a single painful genital ulcer, with associated inguinal lymphadenopathy occurring 2 to 7 days following exposure (34–36). Keratinocytes are likely the first cell type encountered by H. ducreyi upon infection of human skin; thus, the interaction between H. ducreyi and keratinocytes is likely important in establishing infection (37). Chancroid occurs most often in developing countries, including much of Asia, Africa, and Latin America. Epidemics of disease are associated with low socioeconomic status, poor hygiene, prostitution, and drug abuse. Commercial sex workers are also believed to serve as reservoirs for H. ducreyi. After 1987, reported cases of chancroid in the United States declined steadily until 2001. Since then, the number of cases reported has fluctuated from 17 to 55 cases annually. In 2010, only 24 cases were reported to the CDC, with 12 of these cases from Texas (38). Because of difficulties in establishing an etiologic diagnosis of H. ducreyi infection and limited resources in many countries of endemicity, the true incidence of chancroid is unknown. The incidence of H. ducreyi in the United States and other industrialized countries also remains unknown due to poor diagnostic techniques and underreporting, likely making chancroid a more important cause of genital ulcers than is currently reported.

Other Haemophilus spp. H. parainfluenzae remains the predominant species colonizing the URT, accounting for fully 75% of the Haemophilus biota in the oral cavity and in the pharynx. Interestingly, H. parainfluenzae does not routinely colonize the nasal cavity. H. parainfluenzae is thought to account for at least some cases of acute otitis media, acute sinusitis, and acute bacterial exacerbation of chronic bronchitis, although its role in these diseases is often inconclusive. Infrequently, it has also been identified as a cause of subacute bacterial endocarditis. As is the case with systemic infections due to NTHi, blood cultures can be falsely negative in patients with H. parainfluenzae endocarditis due to the fastidious nature of the pathogen and the potential lysis of the organism in the high concentrations of sodium polyanethol sulfonate present in blood culture bottles (39–41). H. aegyptius, a distinct species of Haemophilus that closely resembles biotype III strains of H. influenzae and that has been referred to as the Koch-Weeks bacillus, is an important cause of acute purulent conjunctivitis (42). This disease, often called pinkeye, occurs most often in younger children, especially those having extensive contact with other children in closed settings, such as day care centers and grammar school classrooms. It is characterized by the rapid onset of conjunctival inflammation, visual disturbance, ocular pain

and pruritus. It often involves both eyes and is highly transmissible. Brazilian purpuric fever, a condition that occurs most often in South America, is characterized by rapid onset of high fevers, hypotension, diffuse cutaneous hemorrhaging, and abrupt vascular compromise (43). The causative agent is often mistaken to be H. aegyptius but is instead an organism that is classified in biogroup III of H. influenzae (43, 44). These strains are characterized by the inability to ferment D-xylose, by a particular pattern of their housekeeping genes, by a distinct rRNA restriction pattern, and by resistance to serum bactericidal activity, making them unique among known H. influenzae biogroups (5). H. haemolyticus has recently been reported as a cause of invasive disease in the United States, mischaracterized as H. influenzae (45). The report by Anderson et al. (45) retrospectively evaluated 161 NTHi isolates collected from 2009 to 2010 and 213 isolates collected from 1991 to 2000 and identified 7 strains associated with bacteremia, septic arthritis, or peritonitis that were mischaracterized as H. influenzae. In each instance, strains were poorly hemolytic or nonhemolytic and characterized by phenotypic microbiological identification techniques as H. influenzae. The report emphasized the importance of molecular identification techniques in characterizing NTHi strains. Other Haemophilus species have only rarely been implicated as causes of infection in humans, although lower respiratory tract infection, sinusitis, conjunctivitis, bacteremia, meningitis, wound infections, peritonitis, arthritis, osteomyelitis, and brain abscess have been documented in individual case reports or small case series (7, 46).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS The collection of specimens for the diagnosis of Haemophilus infections is predicated on the nature of the infection being evaluated. Details of specimen collection and transport can be found in chapter 18. In patients suspected of having meningitis, blood and CSF cultures should be performed. Middle ear fluid obtained by tympanocentesis is the specimen of choice for patients with otitis media; however, in patients with perforated tympanic membranes and otorrhea, an aseptically collected aspirate of middle ear fluid from the external auditory canal is also satisfactory. In cases of maxillary sinusitis, direct sinus aspirates or middle meatal swab specimens collected under endoscopic guidance should be obtained. Conjunctival swab specimens are required in the evaluation of patients thought to have Haemophilus conjunctivitis. In patients suspected of having bronchopulmonary infections due to Haemophilus spp., specimens representative of lower respiratory tract secretions should be obtained in such a way as to avoid contamination with oropharyngeal commensal biota. This means that collection of optimal specimens, such as by bronchoalveolar lavage or bronchial washing (less preferable), should be performed to provide optimum specificity when evaluating patients suspected of having Haemophilus bronchopulmonary infections. While collection of sputum and tracheal aspirates is less invasive, distinguishing between pathogens and oral biota can be nearly impossible by this means. When bacterial pneumonia is suspected, blood cultures should also be obtained. Importantly, with one exception, nasal, nasopharyngeal, and nasal swab specimens are of no value whatsoever in evaluating patients suspected of having Haemophilus infections at any of these respiratory tract sites. The one possible exception is in cystic fibrosis patients experiencing

36. Haemophilus n 671

an exacerbation. In this setting, an induced deep-cough specimen collected on a swab inserted into the posterior pharynx may be rewarding (47). In patients suspected of having Haemophilus infections in normally sterile sites, such as the pleural space, synovium, pericardium, or peritoneum, fluid aspirated aseptically from the site of involvement represents the specimen of choice. Concomitant blood cultures should also be performed. Finally, specimens for culture of H. ducreyi should be collected from the margins of genital lesions with a salineor broth-moistened swab. The swab should be immediately transported to the laboratory and plated without delay to avoid loss of organism viability. It is imperative that health care providers inform the laboratory of the clinical suspicion of chancroid so that appropriate media for culture of H. ducreyi can be employed. If extended transport is required, swab specimens should be plated directly at the time of collection in the patient care area or the specimen swab should be placed in transport medium containing hemin (48). When refrigerated (4°C), the use of Amies transport medium has been demonstrated to maintain the viability of H. ducreyi for up to 3 days. Alternatively, specially formulated thioglycolate-hemin-based media containing albumin and glutamine can also be used to preserve organism viability for transport taking >3 days (35, 49). Studies evaluating the viability of H. ducreyi using flocked swabs and liquid Amies transport media are limited, and additional studies are needed. While optimal cultivation of H. ducreyi is based on collection of ulcer materials, lymph node aspirates, pus, and aspirates from buboes can also be submitted for culture, albeit with less sensitivity than ulcer material. When cultivating H. ducreyi from these specimens, laboratories should consider allowing clinicians to directly plate specimens to maintain optimal recovery. Specimens for H. ducreyi nucleic acid amplification techniques should be collected using standard collection techniques for nucleic acid amplification from genital specimens, although large-scale studies evaluating specific transport media have not been conducted. Long-term storage of Haemophilus spp. is usually accomplished by lyophilization or freezing of isolates at −60 to −80°C in tryptic soy broth with >10% glycerol or on porous beads (Pro-Lab Diagnostics, Round Rock, TX).

FIGURE 2 Gram stain of Haemophilus influenzae present in CSF. doi:10.1128/9781555817381.ch36.f2

with Haemophilus spp. In rare cases, the CSF Gram stain may reveal many polymorphonuclear leukocytes but no bacteria. When this occurs, prepare another cytospin smear and stain with acridine orange (BD, Sparks, MD; or Remel, Lenexa, KS). The same approach as that applied to Gram staining CSF should be applied to pleural, peritoneal, synovial, and pericardial fluid specimens. Middle ear fluid specimens and sinus aspirate Gram stains should be prepared directly from the specimen without cytocentrifugation. Gram stains of lower respiratory tract secretions may be prepared directly from the specimen (e.g., expectorated sputa, endotracheal suction specimens, transbronchial biopsy specimens, bronchial brush biopsy specimens, and thoracotomy specimens) or following cytocentrifugation (e.g., bronchial washes and bronchoalveolar lavage fluid). Strands of small, Gram-negative bacilli arranged in a railroad track-like manner on a direct Gram stain are highly suggestive of H. ducreyi. However, Gram staining of genital specimens for H. ducreyi is controversial because most genital ulcers contain a mixed bacterial biota, making Gram stain interpretation difficult. Furthermore, the positive yield of a Gram stain for H. ducreyi is low in comparison to that of culture or detection with nucleic acid amplification tests.

DIRECT EXAMINATION Antigen Detection Microscopy On Gram stain, Haemophilus spp. appear as small, pleomorphic, Gram-negative coccobacilli with coccoid, coccobacillary, rod-shaped, or filamentous forms (Fig. 2). Because of the pleomorphism of Haemophilus spp., careful interpretation of the Gram stain smears must be undertaken to avoid confusion with other Gram-negative bacteria, such as Neisseria meningitidis. Underdecolorization of Gram stains may erroneously suggest the presence of Streptococcus pneumoniae, Listeria monocytogenes, or Streptococcus agalactiae. Gram stain smears of CSF should be prepared and examined and the results reported within 1 h of receipt of the specimen in the laboratory directly to the health care provider who requested the test. With nonturbid specimens, following centrifugation in a cytocentrifuge at 10,000 × g for 10 min, a concentrated smear is prepared. With visibly turbid specimens, a direct smear should be prepared in addition to the cytospin smear. A Gram stain is performed immediately and examined for the presence of polymorphonuclear leukocytes and bacteria morphologically compatible

Commercial immunochemical techniques are available for the detection of S. pneumoniae, Streptococcus dysgalactiae, H. influenzae, and N. meningitidis directly from CSF and other body fluids. While these techniques provide a rapid identification of the pathogen, they lack sensitivity and specificity compared with Gram staining (50). Thus, the use of H. influenzae antigen detection is of limited clinical value and is generally discouraged. However, in certain clinical contexts, such as in resource-constrained regions when the prevalence of disease is high and routine culture is unreliable, antigen-based detection methods may prove useful.

Molecular Techniques Nucleic acid amplification assays, most notably assays predicated on PCR, have been developed to detect H. influenzae directly in various clinical specimens, including CSF, plasma, serum, and whole blood (27, 51). These techniques can be multiplexed to detect other common bacterial causes of specific infectious disease entities, such as meningitis.

672 n BACTERIOLOGY

While publications cite variable detection sensitivities of these techniques, specificity is generally excellent (52–54). Studies have demonstrated that the diagnostic accuracy of chancroid based on clinical presentation and patient history ranges from 33 to 80% (55) and that culture is ∼75% sensitive (56, 57), making it an ideal target for molecular techniques. Molecular strategies have been developed to directly detect H. ducreyi from clinical specimens. Primers for these assays have been designed to amplify sequences from either the H. ducreyi 16S rRNA gene, the rrs (16S)rrl (23S) ribosomal intergenic spacer region, an anonymous fragment of cloned H. ducreyi DNA, or the groEL gene, which encodes the H. ducreyi heat shock protein (56). One strategy includes a chloroform extraction followed by a onetube nested PCR directed to the 16S rRNA gene, with longer outer primers for annealing at a higher temperature and shorter inner primers labeled with biotin and digoxigenin for binding with streptavidin and colorimetric detection (58). Another strategy is to target the hhdA gene; a protocol utilizing this target has been described by Chen and Ballard (59). The sensitivity of PCR directly from clinical specimens varies among assays between 83 and 95% compared to culture or clinical diagnosis (56, 58). The adaptations of molecular methods offer superior sensitivity for the diagnosis of chancroid; they are clearly advantageous in areas where the organism is endemic, particularly where testing by culture is difficult or impossible. In addition to direct detection of H. ducreyi from clinical specimens, studies have also applied PCR to direct detection of H. influenzae and H. parainfluenzae from clinical specimens. In one study by Kuhn et al. (60), the authors evaluated three commercial PCR assays for detection of infectious agents associated with infective endocarditis. This study demonstrated superior performance of PCR compared with culture using both whole blood and valve tissue; the sensitivity of PCR reached 85%, compared with 45% for culture from both whole blood and heart valve tissue. However, while this study resulted in high sensitivity for PCR, the specificity of the PCR was low (40%) due to culture-negative specimens, likely as a result of antimicrobial treatment of patients prior to collection of diagnostic specimens (60). In a second study on meningitis, Wu et al. (61) demonstrated that the sensitivity of real-time PCR and Gram stain was less affected by the presence of antibiotics than culture and Gram stain for detection of H. influenzae, N. meningitidis, and S. pneumoniae. The study further demonstrated that the sensitivity of PCR and Gram stain exceeded that of culture (95.7, 98.25, and 81.3%, respectively) and resulted in identification of an additional 33 (of 451 total specimens) cases of meningitis that were culture negative (61). Other studies have evaluated molecular methods to detect Haemophilus spp. from respiratory specimens (62–64), demonstrating sensitivity of up to 95%, but did not correlate detection of H. influenzae with clinical disease. The use of molecular methods for the identification of other Haemophilus spp. directly from clinical samples has proven difficult. The lack of both sensitivity and specificity has been problematic. In clinical specimens, small numbers of organisms may be present, leading to limitations in detection sensitivity. That is especially the case in patients with Haemophilus bacteremia (65). To achieve adequate sensitivity, large volumes of blood or CSF must be processed, creating laborious nucleic acid extraction and concentration processes with little clinical relevance. Another complicating factor is related to the influence of antimicrobial therapy. While culture frequently becomes negative following administration of the first appropriate dose of antimicrobials,

patients can retain bacterial DNA in their blood or CSF for at least 2 weeks following clearance of the organism, leading to false-positive reactions, often making the interpretation of results challenging. In certain other specimens, particularly specimens from the respiratory tract, commensal strains of Haemophilus spp. are often present, rendering positive results inconclusive. Further, the presence of other commensals or use of antimicrobials prior to screening may result in false-positive reactions, thus contributing to a lack of assay specificity. For these reasons, the use of molecular detection techniques is not currently advocated for the detection of Haemophilus spp. directly in clinical specimens until clinically significant thresholds for molecular quantification of organisms from respiratory specimens are achieved.

ISOLATION PROCEDURES Media Optimum recovery of Haemophilus spp. in culture requires the use of enriched media that support the growth of these fastidious bacteria. Media must contain at least 10 μg/ml of free X and V factors. High concentrations of both the X and the V factor are found in whole blood, most of it sequestered within erythrocytes. X factor is protoporphyrin IX and can be derived from whole blood or can be added to bacteriological media using crystalline hemin; X factor is readily available in standard blood agar. V factor is composed of nicotinamide complexed as NAD or NADP and is also readily available in blood. However, nicotinamide is not readily bioavailable because of its intracellular location and the presence of NAD-glycohydrolase enzymes in blood. For the growth of Haemophilus spp. on solid media, either crystalline hemin and NAD must be added to a final concentration of 10 μg/ml, or the blood used in the medium must be heated such that the red cells lyse and release free X factor and V factor into the medium. The latter can be accomplished by adding blood to the basal medium as it cools to 80°C after being autoclaved. This is referred to as “chocolatizing” blood. For optimal growth of Haemophilus spp., a concentration of 5% chocolatized sheep blood should be employed (1). The optimum growth of Haemophilus spp., especially of more-fastidious species, such as H. ducreyi and H. aegyptius, requires, in addition to the X and V factors, supplementation of media with various other growth factors. Two commercially available supplements that supply these growth factor requirements are IsoVitaleX (BD) and Vitox (Remel). These growth factors contain glucose, cystine, glutamine, adenine, thiamine, vitamin B12, guanine, iron, and aminobenzoic acid and provide adequate supplementation for the growth of H. ducreyi and H. aegyptius. Enriched chocolate agar containing 5% lysed sheep red blood cells and supplemented with 1% IsoVitaleX or 1% Vitox represents one general-purpose medium that is commonly used in clinical laboratories to effectively propagate Haemophilus spp. (Fig. 3). Another medium that reliably supports the growth of Haemophilus spp. is Levinthal medium (66). Because of its transparency, Levinthal medium offers the added benefit of permitting the detection of colony iridescence, a property that is frequently associated with encapsulation (Fig. 4) (67, 68). One investigation found that a medium consisting of GC (BD or Remel) agar base, 5% heated sheep red blood cells, and 1% yeast autolysate provided the best growth of all Haemophilus spp. other than H. ducreyi (69).

36. Haemophilus

n

673

FIGURE 3 Colony morphologies of type b encapsulated (left) and nonencapsulated (right) strains of Haemophilus influenzae when propagated on enriched chocolate agar. doi:10.1128/9781555817381.ch36.f3

A significant challenge in the recovery of Haemophilus spp. from respiratory tract specimens is bacterial overgrowth due to the presence of other less fastidious commensal bacteria. Supplementation of media with some combination of bacitracin, vancomycin, and/or clindamycin serves to inhibit overgrowth with commensals, thus permitting the recovery of Haemophilus spp. (70). Use of such selective media is particularly relevant to the recovery of Haemophilus spp. from respiratory tract specimens from patients with cystic fibrosis, acute exacerbation of chronic bronchitis, conjunctivitis, and epiglottitis (Fig. 5) (71). Several versions of selective Haemophilus media are available commercially, including Haemophilus isolation agar (Remel) and Haemophilus isolation agar with bacitracin (BD). These media contain beef heart infusion agar with casein peptone to supply nutritional requirements, combined with horse blood to supply the X and V factors and to distinguish hemolytic species

FIGURE 4 Colony morphologies of type b encapsulated (left and right) and nonencapsulated (center) strains of Haemophilus influenzae when propagated on Levinthal agar. Note the conspicuous iridescence apparent with the encapsulated strain. doi:10.1128/9781555817381.ch36.f4

of Haemophilus from those that are not. Bacitracin is added to inhibit normal biotas, including Neisseria spp. The use of selective media is also helpful in recovering H. ducreyi from genital tract specimens (49). Selective media may include any of the following: GC agar base with 1% IsoVitaleX, 5% fetal bovine serum, 1% hemoglobin, and 3 μg vancomycin; GC agar base with 5% Fildes enrichment, 5% horse blood, and 3 μg vancomycin; 5% fresh rabbit blood agar with 3 μg vancomycin; or Mueller-Hinton agar with 5% chocolatized horse blood, 1% IsoVitaleX, and 3 μg vancomycin (49). Preferably, two different selective media are employed to ensure optimal sensitivity (49). Growth of Haemophilus spp. may also be achieved on 5% sheep blood agar by use of the microsatellite phenomenon, although it is not recommended for routine clinical testing. With the microsatellite test, a single streak line of hemolysinproducing Staphylococcus spp. is placed on an agar surface previously inoculated with a specimen suspected of containing Haemophilus spp. The hemolysin produced by the Staphylococcus species lyses the erythrocytes immediately adjacent to the streak line in the medium, releasing sufficient concentrations of X factor (hemin) and V factor (NAD) into the medium to supply the growth factor requirements of Haemophilus spp. Staphylococcus also secretes NAD into the medium in proximity to the streak line. Colonies of Haemophilus thus appear in a narrow zone adjacent to the staphylococcal streak. This is referred to as “satelliting” growth (Fig. 6). Organisms other than staphylococci can also produce the satellite phenomenon with Haemophilus, e.g., enterococci and yeast. Although Haemophilus spp. are not a common cause of bacteremia, special techniques are not necessary for their recovery from blood specimens with modern, continuously monitoring blood culture systems (7, 72). The broth medium used in such systems supports the growth of Haemophilus spp. because the blood specimen itself supplies adequate concentrations of both the X and V factors when the erythrocytes present in the specimen lyse as they come into contact with the blood culture broth. However, the common practice of using such systems for the culture of normally sterile body fluids, e.g., synovial, peritoneal, pericardial, and pleural fluid, may be problematic insofar as these specimens may not contain sufficient amounts of blood to supply the necessary levels of the X and V factors to support the growth of Haemophilus spp. In situations where Haemophilus spp. is strongly suspected in such specimens, blood culture bottles

674 n

BACTERIOLOGY

FIGURE 5 Colony growth from an expectorated-sputum specimen containing Haemophilus influenzae from a patient with cystic fibrosis propagated on enriched chocolate agar (left) and enriched chocolate agar containing bacitracin, clindamycin, and vancomycin (right). doi:10.1128/9781555817381.ch36.f5

should be supplemented with at least 10 μg of both sterile hemin and NAD/ml prior to inoculation with clinical specimens. The use of commercial supplements such as Fildes enrichment (BD, Franklin Lakes, NJ) can also be used to aid in the propagation of Haemophilus spp. Following inoculation, solid media should be incubated at 35 to 37°C in a moist atmosphere and in the presence of 5 to 7% CO2. Under these conditions, most Haemophilus spp. grow within 24 to 48 h. When specimens for H. ducreyi and H. aegyptius are cultured, incubation may be necessary for up to 5 days to allow sufficient time for the growth of these fastidious organisms. Further, when technologists attempt to propagate H. ducreyi, plates should be incubated at slightly lower temperatures, i.e., 30 to 33°C in 5% CO2 in a high-moisture environment. Use of lower incubation temperatures will improve the recovery of H. ducreyi in comparison to incubation temperatures of 35 to 37°C.

Colony Appearance Colonies of Haemophilus spp. on suitable solid media, in general, are nonpigmented or slightly yellow and flat to

FIGURE 6 Satellite phenomenon observed when Haemophilus influenzae is propagated next to a streak of Staphylococcus aureus on a 5% sheep blood agar plate. doi:10.1128/9781555817381.ch36.f6

convex, and have a diameter of 0.5 to 2 mm after 48 h of incubation. Certain species of Haemophilus produce betahemolysis (Table 1). Colonies of H. influenzae on chocolate agar are smooth, low, convex, grayish, and translucent. Encapsulated strains often have a mucoid appearance, while nonencapsulated strains produce smaller, buff colonies (Fig. 3). Most strains of H. influenzae produce indole, emitting a strong amine-like odor. Non-indole-producing strains emit a “mousy” odor. Colonies are 1 to 2 mm in diameter and often grow within 24 h. Colonies grown on clear media, such as Levinthal agar, demonstrate iridescence under obliquely transmitted light (67, 68). Iridescence is most conspicuous with young colonies and disappears with age. Iridescent colors may include yellow, red, green, or blue. Iridescence is more apparent with capsular type b strains; nonencapsulated strains typically demonstrate a blue-green color (Fig. 4). Colonies of H. aegyptius reach a colony size of only ca. 0.5 mm after 48 h of growth. Colonies are low, convex, and translucent, with a smooth, entire surface. On semisolid agar media (which contains ∼0.4% agar, compared with 1.5% for solid media), “comet-like” colonies are produced (5, 42). Colonies of H. parainfluenzae are typically off-white to yellow and, like H. influenzae, 1 to 2 mm in diameter after 24 h of growth. The colony appearance is extremely varied, i.e., flat and smooth, granular with serrated edges, or heaped up and wrinkled. Colonies exhibiting the last morphology may be slid intact across the surface of the agar. The colony morphology of H. parainfluenzae may change as the colonies age. Colonies of H. haemolyticus are translucent, smooth, and convex and do not form satellites around Staphylococcus. Colonies usually achieve a diameter of 0.5 to 1.5 mm after 24 h, with a clear zone of beta-hemolysis surrounding each colony when the organism is grown on blood agar; H. haemolyticus can lose its ability to cause hemolysis following serial subculture on bacteriological media. The growth properties and colony morphology of H. parahaemolyticus and H. paraphrohaemolyticus are similar to those of H. haemolyticus. H. ducreyi grows poorly, regardless of the medium used, and frequently 3 to 5 days will pass before growth appears. Colonies growing on chocolate agar are small, flat, gray,

36. Haemophilus n 675

and smooth. Larger colonies may be interspersed among small colonies but have the same morphology. Growth on blood agar is poor, with a slight beta-hemolysis surrounding the colonies. As with H. parainfluenzae, older colonies of H. ducreyi are cohesive and can be slid across the agar.

IDENTIFICATION The identification and differentiation of Haemophilus spp. are achieved through determination of X and V factor requirements for growth; performance of the porphyrin test; assessment of hemolysis; determination of carbohydrate fermentation patterns; and production of indole, ornithine decarboxylase, urease, catalase, and β-galactosidase (Table 1). The pattern of X and V growth factor requirements and the porphyrin test provide sufficient information for the presumptive species identification of selected Haemophilus spp. Definitive species identification, however, requires assessment of the other phenotypic characteristics listed above (Table 1). Alternatively, sequencing of the 16s rRNA gene or matrix-assisted laser desorption ionization–time-of-flight mass spectrometry (MALDI-TOF MS) also provide definitive identification of these organisms.

X and V Factor Growth Requirements X and V factor requirements for the growth of Haemophilus spp. may be determined by swab inoculation of a suspension of test organism equivalent in turbidity to a 0.5 McFarland standard across the entire surface of a 100-mm petri dish containing tryptic soy agar. Filter paper disks or strips impregnated with X factor, V factor, and the X and V factors (Remel or BD) are then placed on the agar surface, and the plate is incubated for 20 to 24 h at 35°C in an atmosphere of 5 to 7% CO2. The pattern of satellite growth around individual disks or strips, in the absence of growth elsewhere on the plate, is used to define the growth factor requirements of the test strain (Fig. 7). Tryptic soy agar is the preferred medium for use when the X and V growth factor requirements for Haemophilus spp. are determined, as other media

FIGURE 7 Use of X and V factor disks and strips in determining the growth factor requirements of Haemophilus influenzae (left disks) and Haemophilus parainfluenzae (right disks). doi:10.1128/9781555817381.ch36.f7

may yield erroneous results (73, 74). Alternatively, triplates (Haemophilus ID II; Remel) and quadplates (Haemophilus ID Quad; Remel) can be used to assess X and V growth factor requirements. When performing X and V factor studies, care should be taken to avoid carrying X factor along with the inoculum. This can result in erroneous identification of H. influenzae as H. parainfluenzae. All of the X factor-requiring species of Haemophilus, most notably H. influenzae, lack the enzymes necessary to convert δ-aminolevulinic acid (ALA) into protoporphyrin, a metabolic intermediate in the biosynthesis of X factor (75). Thus, they require that X factor be supplied exogenously in order to support growth. By taking advantage of this observation, a rapid test, known as the porphyrin or ALA test, can be performed to quickly determine if a test organism requires X factor for growth (76). When positive, the porphyrin test indicates that the test organism is X factor independent; when negative, the porphyrin test indicates that the organism requires X factor. Since the vast majority of X factorrequiring Haemophilus spp. recovered in the clinical laboratory are H. influenzae, when the porphyrin test is performed with a clinical isolate and found to be negative, it can be inferred with a high likelihood that the organism is H. influenzae. The porphyrin test is performed using commercially available ALA disks (Remel) or through preparation of liquid porphyrin medium (76). To prepare liquid porphyrin medium, 2 mM ALA and 0.8 mM MgSO4 in 0.1 M phosphate buffer (pH 6.9) are aliquoted in glass tubes with 0.5 ml of porphyrin medium in each tube. Tubes can be stored at 4°C for several months or for years at −20°C. Tubes are inoculated with a loopful of freshly grown bacteria and incubated for 4 h at 35°C in ambient air (1). Following incubation, the tubes are examined for brick-red fluorescence with a device, such as a Wood’s lamp, that emits a 360-nm-long-wave UV light (Fig. 8). Tubes with questionable results may be reincubated for up to 24 h. Alternatively, Kovács reagent (0.5 ml) can be added to the liquid porphyrin medium, and the tube can be shaken and observed for a red color in the lower water phase (1). A negative-control tube lacking ALA should also be inoculated when the porphyrin test is performed to rule out falsepositive reactions due to the presence of indole. The porphyrin test has been shown in several studies to outperform

FIGURE 8 Positive (top) and negative (bottom) porphyrin tests. doi:10.1128/9781555817381.ch36.f8

676 n BACTERIOLOGY

growth factor-based methods for differentiation of H. influenzae from non-H. influenzae species (77).

Conventional Biochemical Tests The list of biochemicals necessary for the differentiation of Haemophilus species is provided in Table 1. Carbohydrate fermentation is determined in phenol red broth containing 1% carbohydrate supplemented with 10-μg/ml NAD and hemin. Following heavy inoculation, fermentation tubes are incubated at 35°C without CO2 for up to 1 week and examined periodically for a red-to-yellow color, indicating a positive reaction. Usually positive reactions become apparent within 24 h; however, H. aegyptius and A. segnis have been noted to demonstrate weak reactions following a 24h incubation (1). Indole production can reliably be determined with most strains of H. influenzae and H. parainfluenzae using a spot indole test (Remel or BD). However, reliable assessment of indole production by other Haemophilus spp. requires use of a solution of 0.1% L-tryptophan in 0.067 M phosphate buffer at pH 6.8. Following inoculation, the suspension is incubated for 4 h at 35°C in ambient air, 0.5 ml of Kovács reagent is added to the tube, and the tube is shaken and examined for the appearance of red color in the upper portion of the tube as an indication of a positive reaction (1, 5) (Fig. 9). Production of ornithine decarboxylase is determined using ornithine decarboxylase medium (see chapter 19) or Moeller medium. A tube containing either medium is heavily inoculated, incubated for 4 to 24 h at 35°C in ambient air, and examined for a purple coloration as an indication that the organism produces ornithine decarboxylase (1, 5) (Fig. 9). Species of Haemophilus positive for ornithine decarboxylase include several biotypes of H. influenzae and H. parainfluenzae. As was the case with indole production, in most instances, a spot urease test (Remel or BD) can be used to reliably detect urease production by most strains of H. influenzae and H. parainfluenzae. Determination of urease production with other species requires the use of urease medium containing 0.1 g KH2PO4, 0.1 g K2HPO4, 0.5 g NaCl, and 0.5 ml phenol red (1:500) dissolved in 100 ml of distilled water. The pH is adjusted to 7.0 with NaOH, and 10.4 ml of a 20% aqueous solution of urea is added. After inoculation, the tube is incubated for 4 h at 35°C in ambient air and examined for a pink-to-red coloration, indicating a positive reaction (1, 5) (Fig. 9).

Commercial Biochemical Identification Systems Several commercial identification systems have been developed to identify Haemophilus spp. These systems employ a

battery of conventional biochemical tests, frequently in a miniaturized form, with results available in shorter time periods than with conventional biochemical tests. The performance characteristics and identification accuracy of these commercial systems are extremely variable (69, 73, 77, 78). The RapID NH system (Remel) contains 11 biochemical reactions in a microwell tray. The reactions used for identification of Haemophilus include the production of urease, indole, ornithine decarboxylase, proline, and γ-glutamyl aminopeptidase; resazurin reduction; glucose and sucrose utilization; nitrate reduction; and phosphate hydrolysis. The kit uses phosphate hydrolysis and nitrate reduction reactions to identify an isolate as belonging to the genus Haemophilus and the remaining reactions to identify the isolate to the species level and to determine the biotype of H. influenzae. Although various results have been reported with the RapID NH system, when used properly, >95% of clinical isolates of H. influenzae should be correctly identified (73). The BBL Crystal Neisseria/Haemophilus ID system (BD) and API NH kit (bioMérieux, Marcy l’Etoile, France) provide miniaturized biochemical identification schemes in a microwell tray, with results available in ≤5 h. The Crystal system employs 29 different growth substrates and is predicated on measuring the substrate conversion chromogenically and fluorogenically after 5 h of incubation. The API NH kit consists of 12 dehydrated substrates and a well to detect penicillinase, and it permits the identification of Haemophilus spp., Neisseria spp., and Moraxella spp. The test is performed by inoculating each well with an organism suspension equivalent to a 4 McFarland turbidity standard prepared from 24-h colony growth and incubating the plate for 2 to 2.25 h at 35°C. In addition to testing for penicillinase production, it tests for the following biochemicals: glucose, fructose, maltose, saccharose, ornithine decarboxylase, urease, lipase, alkaline phosphatase, β-galactosidase, proline arylamidase, γ-glutamyl aminotransferase, and indole (79, 80). Independent studies evaluating the performances of the Crystal Neisseria/Haemophilus and API NH kits compared to accepted gold standards have not been published. One instrument-based identification system has been developed for the species identification of Haemophilus spp., the Neisseria-Haemophilus identification cards for use with the VITEK 2 instrument (bioMérieux). This system is based on colorimetric detection of preformed enzyme complexes using chromogenic substrates. The database supporting these cards encompasses 27 taxa, including Neisseria, Haemophilus, Actinobacillus, Campylobacter, Capnocytophaga, Cardiobacterium, Eikenella, Gardnerella, Kingella, Moraxella, Oligella, and Suttonella species. Studies with the VITEK 2 system using both collections of well-characterized stock strains and clinical isolates have demonstrated identification accuracies of 90 to 95%, with results varying by species (78, 81). In one recent study, >95% of isolates of H. influenzae, A. segnis, H. parahaemolyticus, H. parainfluenzae, and A. actinomycetemcomitans were correctly identified, while none of the test strains of H. haemolyticus were correctly identified (81).

Mass Spectrometry

FIGURE 9 Conventional biochemicals depicting positive and negative reactions for indole, urease, and ornithine decarboxylase production (left to right) by Haemophilus spp. doi:10.1128/9781555817381.ch36.f9

Studies evaluating MALDI-TOF MS largely demonstrate excellent performance for HACEK group organisms (i.e., Haemophilus spp., Aggregatibacter spp., Cardiobacterium spp., Eikenella corrodens, and Kingella spp.), including Haemophilus species (82, 83). Couturier et al. (82) demonstrated that >90 and >98% of H. parainfluenzae and H. influenzae strains, respectively, could be identified to the species level using MALDI-TOF. Similarly, van Veen et al. (83) demonstrated >98% identification of HACEK organisms to the species

36. Haemophilus

level. Branda et al. (84) recently evaluated the VITEK MS IVD (bioMérieux) and found 96 and 92% correct identification to the species level for H. influenzae and H. parainfluenzae, respectively. Comparisons of the Bruker MALDI Biotyper (Bruker Daltonics, Billerica, MA) to the VITEK MS are limited; however, Frickmann et al. (85) compared the VITEK MS RUO (bioMérieux) with both the Biotyper and fluorescence in situ hybridization for identification of H. influenzae, H. parainfluenzae, and H. parahaemolyticus. With formic acid extraction, the Biotyper identified 100% of H. influenzae (50/50), 88% of H. parainfluenzae (22/25), and 100% of H. parahaemolyticus (2/2) strains to the species level. Results were not significantly different for the Biotyper without prior formic acid extraction, although identification of H. parainfluenzae dropped to 72% (18/25) and the Biotyper missed an additional H. parahaemolyticus identification. The VITEK MS RUO identified 82% of H. influenzae (41/50), 68% of H. parainfluenzae (17/25), and 50% of H. parahaemolyticus (1/2) strains. All identifications on the VITEK MS were performed without prior formic acid extraction (85). Neither system identified any of the seven H. haemolyticus strains analyzed (85), likely due to poor representation of the organism in the databases. Martiny et al. (86) compared the performance of the VITEK MS IVD to the Biotyper IVD and found 100% identification (n = 30) of H. influenzae by both systems. Evaluation of species other than H. influenzae in this study was limited by the lack of diversity of species included in the study (86). Performance of MALDI-TOF on Haemophilus strains other than those discussed above and performance of MALDI-TOF for typing H. influenzae have not been evaluated, and further study is warranted.

Molecular Identification Several molecular methods, including 16S rRNA gene sequencing, next-generation sequencing, PCR, microarrays, and fluorescence in situ hybridization, have been described in the literature as being effective tools for the species identification of Haemophilus spp. when performed on organisms recovered in culture (51, 87–89). Molecular targets for the detection and identification of Haemophilus spp. are numerous. Previous studies have described the detection of H. influenzae using the cap locus (which includes the capsule bexA) (53, 63, 90, 91), the 16S rRNA gene (51, 63, 92, 93), the insertion-like sequence (IS1016) (91), the fumarate reductase iron-sulfur gene B (frdB) (94), the manganese-dependent superoxide dismutase (sodA) (95), and the outer membrane protein P6 gene (ompP6) (63, 92, 96). Many of these targets are then combined with PCR (real time or traditional), microarrays, or sequencing to identify the organism. Widely utilized for sequencing, 16S rRNA frequently resolves the identities of strains to the species level; however, identification of H. influenzae, H. aegyptius, and H. influenzae biogroup aegyptius can be problematic due to the high degree of homology in their sequences. In these instances, sequencing of other targets, such as ropD, should be considered, or the use of combined sequencing and biochemical studies can be used. Application of highly multiplex disease-state panels such as the FilmArray (BioFire Diagnostics, Salt Lake City, UT) or Verigene (Nanosphere, Inc., Northbrook, IL) to positive blood cultures has demonstrated reduction in length of stay and enhanced antimicrobial stewardship (97). Application of these technologies to identification of Haemophilus spp. from positive blood cultures may offer a significant reduction in turnaround time; however, peer-reviewed studies evaluat-

n

677

ing these technologies in Haemophilus have not been published. Next-generation sequencing has been applied to Haemophilus spp. in two recent studies without the need for prior culture. In a study by Salipante et al. (98), the authors conducted deep sequencing on 66 sputum specimens from cystic fibrosis patients and identified H. influenzae in 5 specimens. Of the 5 specimens that yielded H. influenzae, 4 could be detected only via sequencing and 1 could be detected only in culture (98). The discrepancy between the results of culture and sequencing illustrates the challenge of correlating cultures with sequencing. While the presence of an organism on culture definitively confirms the presence of a viable organism, the presence of DNA detected in a sequencing reaction may still represent an infectious process but can also represent a remnant of a previous infection. Attempts to resolve this dilemma are important since, at a cost of less than $75 in reagents per sample, sequencing has quickly become affordable for the routine clinical microbiology laboratory (98).

Problems in Identification A significant challenge in the species identification of H. influenzae, H. aegyptius, and H. influenzae biogroup aegyptius is a lack of biochemical diversity and sequence divergence (1, 5, 7). Biochemical profiling and standard 16S rRNA gene sequencing fail to adequately distinguish these organisms (1, 5, 7), necessitating the use of alternate sequencing targets, such as those mentioned in the previous section, or a combination of sequencing and biochemical testing. This is problematic, since H. aegyptius lacks the potential to cause Brazilian purpuric fever, while strains of H. influenzae biogroup aegyptius cause the disease. Xylose fermentation by most H. influenzae isolates combined with the lower growth rate of H. influenzae biotype aegyptius and the ability of H. aegyptius to agglutinate human erythrocytes may be of some value in distinguishing these organisms (1, 7, 99).

TYPING SYSTEMS Capsular Serotyping and Biotyping The capsular antigen of H. influenzae is a principal virulence determinant of this organism. Six different capsular antigens have been recognized, each of which is characterized by a distinct carbohydrate chemical composition and given a letter designation from “a” to “f.” Prior to the introduction of the pediatric HIB vaccine, capsular type b strains were recovered from human clinical material most often. However, today, at least among populations in which there is widespread use of the HIB vaccine, non-b encapsulated strains occur with nearly equal frequency. For this reason, it may be instructive to know the capsular serotypes of H. influenzae strains recovered from clinical specimens, especially those representative of invasive disease. This may be accomplished using both phenotypic and genotypic methods. Capsular serotyping of H. influenzae is best accomplished by the use of a slide agglutination assay that employs polyclonal antisera specifically reactive with each of the six capsular antigens (5, 100, 101). It is advisable to perform serotyping as soon as possible after isolation of H. influenzae, as the amount of capsular antigen produced may diminish over time, especially with repeated subculture. A thick, homogenous suspension of test organism is prepared in saline, 1 to 2 drops are placed on a glass slide, and then a drop of type-specific antisera is added. The antisera is mixed with the organism suspension, and then the glass slide is rocked gently for ca. 1

678 n BACTERIOLOGY

min before being examined for the presence of clumping, an indication of a positive reaction. The reagents for performing slide agglutination serotyping of H. influenzae are commercially available in kit form from Remel and BD. Alternatively, primary type-specific antibodies can be directly or indirectly detected with fluorescent molecules, and binding of the antibody to the homologous capsular antigen can be determined by fluorescence microscopy (102, 103). While these technologies remain viable, reagents are not commercially available. Whether a slide agglutination test or fluorescent antibodies are used to determine the capsular serotype of an isolate of H. influenzae, positive- and negative-control strains should always be processed simultaneously with clinical isolates as a means of validating test results. As noted above, based on three phenotypic properties, the production of indole, ornithine decarboxylase, and urease, strains of H. influenzae and H. parainfluenzae can be distinguished into multiple different biotypes (Table 2). Also, as outlined previously, at least with H. influenzae, certain biotypes have been found to have specific disease associations. Assessment of indole, ornithine decarboxylase, and urease production with clinical isolates of H. influenzae and H. parainfluenzae can be accomplished using the conventional methods described above (Fig. 9) or by use of commercially available miniaturized biochemical kit systems (11). In one recent study that compared the API NH strip kit (bioMérieux) with the RapID NH system (Remel) and the Neisseria-Haemophilus identification card (bioMérieux) as a means for determining the biotypes of a large collection of recent clinical isolates of both H. influenzae and H. parainfluenzae, the API NH kit yielded the most reliable results, correctly classifying the biotypes of >97% of the strains tested (80).

Typing by Molecular Methods Molecular methods have the advantage of enhanced sensitivity and specificity (104) due to the use of standardized techniques and a lack of false-positive reactions observed with nonencapsulated strains in slide agglutination tests. Capsular typing of H. influenzae can also be accomplished by use of various molecular methods. Most such assays rely on the amplification of genes in the cap locus, the outer membrane protein D gene (glpQ), the capsule-producing gene (bexA), the 16S rRNA gene, and the insertion-like sequence (105). One algorithm was used for detection of the cap genes to determine capsular serotypes a through f, while the capsuleproducing gene, bexA, was used to separate strains that produce capsule from those that do not (105). Detection of the ompP2 (outer membrane lipoprotein P2) gene was used as a control. Using this system, both a conventional PCR and a real-time PCR assay were found to be more sensitive than a slide agglutination test for serotyping H. influenzae (105). Similarly, Wroblewski et al. (106) described a two-step algorithm combining a two-plex real-time PCR with a five-plex serotype-specific real-time PCR (type a, acsA; type c, ccsB; type d, dcsC; type e, ecsC; and type f, fcsA). In this algorithm, the two-plex PCR containing primers targeting bexA (capsule transport gene) and bcsB (type b-specific gene) is used to screen isolates for NTHi and serotype b-specific strains. Strains that are positive for bexA but negative for bcsB are referred to the five-plex PCR for serotype-specific amplification (106). This method demonstated 100% concordance with slide agglutination (106). As with the Enterobacteriaceae, pulsed-field gel electrophoresis (PFGE) is considered the gold standard for strain typing of Haemophilus. The method demonstrates excellent

separation of clones but is laborious and time-consuming (107–109). Other molecular methods for typing have also been applied to Haemophilus species. Studies evaluating repetitive-element sequence-based PCR using intergenic dyad sequence (IDS)-specific primers (IDS-PCR) for nonencapsulated Haemophilus strains have been developed and demonstrate excellent separation of NTHi strains (108). In one study evaluating the performance of IDS-PCR with 69 NTHi isolates, the assay demonstrated 65 different banding patterns that were epidemiologically classified as fingerprints similar to those obtained by PFGE (108). Other typing technologies applied to Haemophilus with a high degree of separation include ribotyping, restriction fragment length polymorphism analysis, multilocus enzyme electrophoresis, randomly amplified polymorphic DNA profile analysis, and multilocus sequence typing (108). While all of these techniques have demonstrated excellent separation, many are laborious and time-consuming (multilocus enzyme electrophoresis, PFGE, and ribotyping), others produce overly complex banding patterns (restriction fragment length polymorphism analyses), and others lack reproducibility (randomly amplified polymorphic DNA profile analysis) (108). Multilocus sequence typing offers the advantage of superior discriminatory power because it combines sequence typing of seven housekeeping genes with results that can readily be compared between laboratories (7). Next-generation sequencing offers the promise of detection of an organism, classification of antimicrobial resistance factors, and typing of the organism in a single assay. Furthermore, with turnaround time for sequencing and annotation of the sequence decreasing to 12 years of age (51, 102–104), so limiting STEC testing to children would miss many infections. Although STEC infections are more common in the summer months, sporadic cases and outbreaks do occur year-round (51, 53, 101). Leukocytes are often but not invariably observed in the stools of patients with STEC infection; thus, determination of white blood cells in stool should not be used as a criterion for STEC specimen selection (51, 105). Enrichment. Although broth enrichment is widely used for the recovery of O157 STEC from foods, there is little evidence that it enhances isolation from human fecal specimens. However, immunomagnetic separation (IMS), a technique shown to increase the rate of isolation of O157 STEC from food specimens, has been adapted to culture of fecal specimens (106). IMS enhances the detection of O157 STEC from patients with HUS, patients presenting an extended period of time after the onset of illness (>5 days), asymptomatic carriers, or specimens that have been stored or transported improperly. IMS beads for O157, O111, and O26 are available commercially (Table 3), or laboratories may produce beads with other O-specific antibodies (107). Plating media. Because O157 STEC strains ferment lactose, they are impossible to differentiate from other lactosefermenting organisms on lactose-containing media. Most O157 STEC strains do not ferment the carbohydrate Dsorbitol overnight, in contrast to the ∼80% of other E. coli strains that ferment sorbitol rapidly. Thus, sorbitolcontaining selective media are often used for isolation of O157 STEC. Sorbitol-nonfermenting colonies are suspected (but not definitively known) to be E. coli O157:H7 (108). In some areas of central Europe, sorbitol-fermenting O157 STEC strains are commonly isolated from patients with HUS (109); these organisms are very rare in North America (Strockbine, unpublished data). Specific culture media have been developed to exploit phenotypic and antibiotic resistance traits that are characteristic of STEC strains. Although sorbitol-containing MAC (SMAC) is widely used, cefixime-tellurite-containing SMAC (CT-SMAC) and CHROMAgar O157 have been shown to increase the sensitivity of culture for O157 STEC (110). It has been reported that some nonmotile strains of O157 STEC fail to grow on CT-SMAC (106). Several chromogenic agar media are available commercially to assist in rapid identification (Table 3); these media generally perform well for O157:H7 and for some non-O157 STEC strains (111–114). Screening procedures for STEC strains. For the isolation of O157 STEC from SMAC, colorless (nonfermenting) colonies are tested with O157 antiserum or latex reagent (115) (Table 3). If the O157 latex reagent is used, it is

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 692

important to test positive colonies with the latex control reagent to rule out nonspecific reactions. The manufacturers of these kits recommend that strains that react with both the antigen-specific and control latex reagents be heated and retested. However, in a study that followed this procedure, none of the nonspecifically reacting strains were subsequently identified as O157 STEC (116). Unlike most other E. coli strains, O157 STEC strains do not express β-glucuronidase; therefore, the MUG reaction (4-methylumbelliferryl-β-D-glucuronide for detection of βglucuronidase activity) is helpful for screening for O157 STEC (117). MUG-positive, urease-positive O157 STEC strains have been isolated in the United States but are still rare (118; Strockbine, unpublished data). For the recovery of STEC strains from stool specimens that test positive for Shiga toxin, CT-SMAC, CHROMagar O157, or a similar selective agar for isolation of O157 STEC should be used. If sorbitol-nonfermenting colonies are negative with O157 latex, then sorbitol-fermenting colonies (because most non-O157 STEC strains ferment sorbitol) and a representative sample of sorbitol-nonfermenting colonies may be selected for Shiga toxin testing. Latex reagents and antisera (Table 3) for detecting certain non-O157 STEC serotypes are now available and could also be used to test colonies from Shiga toxin-positive specimens or to serogroup Shiga toxin-positive isolates. Virtually all O157 STEC strains and 60 to 80% of nonO157 STEC strains produce a characteristic E. coli hemolysin, referred to as enterohemolysin (Ehly), which is distinct from the alpha-hemolysin produced by other E. coli strains (119). Washed sheep blood agar supplemented with calcium (WSBA-Ca) is used as a differential medium for the detection of enterohemolytic activity (119). Ehly-producing colonies can be differentiated from alpha-hemolysin-producing colonies on WSBA-Ca because the latter are visible after 3 to 4 h of incubation. After 3 to 4 h, colonies are marked for the appearance of alpha-hemolysin, and the plates are examined again after 18 to 24 h. Incorporation of mitomycin C into WSBA-Ca enhances the appearance of the Ehly hemolysis and increases the proportion of non-O157 STEC strains that exhibit this activity (120). Because some nonO157 STEC strains do not demonstrate the enterohemolytic phenotype, nonhemolytic colonies should also be screened for Shiga toxin (121). Presumptive STEC isolates should be sent to a reference laboratory or a public health laboratory for further characterization.

Isolation Procedures for Other Diarrheagenic E. coli Methods for the isolation of ETEC, EPEC, EIEC, EAEC, and putative diarrheagenic E. coli strains are generally available only in reference or research settings. Public health and reference laboratories usually examine specimens for these pathogens only when an outbreak has occurred and specimens are negative for routine bacterial pathogens. ETEC and EAEC should be considered possible etiologic agents of watery diarrhea when no other pathogen has been identified, especially for travelers. EPEC should be considered a possible pathogen in outbreaks of severe nonbloody diarrhea occurring in infants or young toddlers, particularly in nursery or day care settings (65). EIEC should be considered a possible etiologic agent in outbreaks of nonwatery diarrhea (bloody or nonbloody). To capture E. coli for further testing, fecal specimens should be plated on a differential medium of low selectivity (e.g., MAC). Five to 20 colonies, mostly lactose fermenting

37. Escherichia, Shigella, and Salmonella n 693

but with a representative sample of nonfermenting colonies, should be selected and inoculated onto nonselective agar slants (such as L agar or nutrient agar). These colonies are then sent to a reference laboratory for testing or are screened for virulence-associated characteristics if assays are available. Strains can be kept frozen for long periods in L broth with 15 to 50% glycerol at −80°C. Arrangements for sending E. coli isolates from well-characterized outbreaks to the CDC for testing can be made through local and state health departments. Screening procedures for ETEC, EPEC, EAEC, and EIEC strains. E. coli pathotypes other than STEC cannot be distinguished from other E. coli strains by phenotypic screening techniques. Many EIEC strains are nonmotile and fail to decarboxylate lysine; however, some EIEC strains are motile or lysine positive. Use of commercial antisera to the classical EPEC somatic (O) and capsular (K) antigens to detect EPEC is no longer recommended due to the number of false positives this approach yields.

Identification Phenotypic Identification With the exception of E. albertii, the commercial identification systems do a good job of identifying most Escherichia strains (122–124). Identification of E. albertii with these systems remains problematic because representative strains of this species are not yet included in commercial databases (125). Abbott and colleagues, who extensively characterized five strains of E. albertii by conventional phenotypic methods and by commercial identification panels, reported that E. albertii is an indole-negative species that ferments Dmannitol but not D-xylose (125). In their study, E. albertii strains were identified by commercial systems as Hafnia alvei, Salmonella or Salmonella enterica serotype Choleraesuis, E. coli (inactive or serotype O157:H7), or Yersinia ruckeri. Although some strains were clearly misidentified, the majority of the strains generated probability scores for the final identification that were unacceptable, or the identification was inconsistent with the source of the specimen (e.g., identification of the fish pathogen Y. ruckeri from a human specimen), which should have triggered additional phenotypic tests to establish a more reliable identification. The authors found that the most reliable clue to the possible presence of E. albertii was an unacceptable first-choice identification of H. alvei for an isolate that is both L-rhamnose and Dxylose negative. Phenotypic tests that can help discriminate E. albertii strains from selected members of the Enterobacteriaceae family with similar phenotypic traits are shown in Table 1. Two biogroups of E. albertii are listed in Table 1. These correlate with two of the distinct clusters of strains identified in the E. albertii lineage by phylogenetic studies (8). Biogroup 1 comprises the five strains isolated from Bangladeshi children with diarrhea, while biogroup 2 comprises strains formerly identified as S. boydii 13. The strains in the two biogroups differ from each other in their abilities to produce indole from tryptophan, decarboxylate lysine, and ferment D-sorbitol. Antigenic relationships between members of the E. albertii lineage and other members of the Enterobacteriaceae family have been observed (e.g., S. boydii 7 and E. coli O28). A diagnostic PCR assay using three housekeeping genes was described by Hyma et al. (8) for E. albertii; this assay is independent of phenotypic or antigenic traits and should facilitate studies to learn about the diversity within the lineage, the natural habitat of the species, and its role in enteric disease.

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 693

The identification of presumptive O157 STEC isolates is necessary because other species may cross-react with O157 antiserum or latex reagents, including Salmonella O group N (O:30), Yersinia enterocolitica serotype O9, Citrobacter freundii, and E. hermannii. Additional phenotypic tests (cellobiose fermentation and growth in the presence of potassium cyanide [KCN]) may be necessary to differentiate E. hermannii from E. coli, but because E. hermannii is rarely detected in stool specimens, use of these tests is not costeffective for most laboratories. In recent years, matrix-assisted laser desorption ionization–time-of-flight mass spectrometry (MALDI-TOF MS) has been adopted in many clinical laboratories for identification. The method is rapid, is substantially less expensive than conventional culture methods, and has performed well in identifying E. coli to the species level (126–129). It holds promise for the identification of other Escherichia species (E. hermannii, E. vulneris, and E. fergusonii) to the species level, but not unexpectedly, it was unable to distinguish Shigella from E. coli (126, 127). Clark et al. (128) recently reported success using custom evaluation criteria (informative spectral peaks) as a screening tool to differentiate most E. coli pathotypes for subsequent testing. In response to the growing problem of multidrug-resistant bacteria, labs are seeking rapid methods to screen bacteria to identify resistance mechanisms to inform and expedite the management of patients to control the spread of resistance. Several groups recently reported MS methods to rapidly test for antimicrobial resistance mechanisms. Jung et al. (130) described a MALDI-TOF MS assay to detect βlactamase activity against aminopenicillins. The assay works by detecting a specific pattern of fragments generated from the digestion of the drug by the resistance-conferring enzyme. The assay took just 2.5 h to complete and had a sensitivity and specificity of 100 and 91.5%, respectively, when evaluated on a set of 100 isolates. Another assay using MALDI-TOF MS to detect class D carbapenemase OXA48-like-producing Enterobacteriaceae was reported by Sauget et al. (131). This assay had a turnaround time of 90 min and a sensitivity and specificity of 98.9 and 97.8%, respectively, when tested on 372 strains. A third assay that uses liquid chromatography-tandem MS to detect carbapenemase KPC, NDM, OXA, IMP, and VIM activity produced against imipenem, meropenem, and ertapenem was described by Kulkarni et al. (132). The sensitivity and specificity of the assay were best with imipenem—96 and 95%, respectively, for a group of 402 Gram-negative bacteria—and the assay took only 75 min to complete. These assays offer valuable information quickly to facilitate interventions to limit the dissemination of resistant organisms and prevent outbreaks. Additional information on the use of MS to detect antibiotic resistance mechanisms is reviewed by Hrabak et al. (133).

Serotyping The serologic classification of E. coli is generally based on the O antigen (somatic) and the H antigen (flagellar) (134). The O and H antigens of E. coli are stable and reliable strain characteristics, and although 181 O antigens and 56 H antigens have been described (a few of which are no longer recognized), the actual number of serotype combinations associated with diarrheal disease is limited (Table 2). Determination of the O and H serotypes of E. coli strains implicated in diarrheal disease is particularly useful in epidemiologic investigations (Table 2). Even though antisera for microtiter or tube agglutination tests are available from manufacturers, most laboratories do not attempt to perform complete E. coli serotyping because it is costly. For well-

694 n BACTERIOLOGY

characterized outbreaks with no identified etiologic agent, arrangements may be made through state health departments to send E. coli isolates to the CDC for virulence testing and serotyping.

Serologic Confirmation of O157 STEC Confirmation of E. coli O157:H7 requires identification of the H7 flagellar antigen. H7-specific antisera and latex reagents are commercially available (Table 3), but detection of the H7 flagellar antigen often requires multiple passages (115). Isolates that are nonmotile or negative for the H7 antigen should be tested for the production of Shiga toxins or the presence of Shiga toxin gene sequences. Approximately 85% of O157 isolates from humans received by the CDC are serotype O157:H7, 12% are nonmotile, and 3% are H types other than H7 (Strockbine, unpublished data). E. coli O157:NM strains often produce Shiga toxin, and those that do produce Shiga toxin have the same fliC restriction profile as that observed for E. coli O157:H7 (135). No O157 strain from human illness with an H type other than H7 has been found to produce Shiga toxin (Strockbine, unpublished data).

Nucleic Acid-Based Methods Accurate identification of bacterial isolates is important for directing patient care and management. Compared to traditional phenotypic approaches, which can be influenced by phenotypic variation or subjective interpretation, 16S rRNA gene sequencing is a more objective identification tool and has the potential to reduce laboratory errors. Some clinical laboratories have begun using molecular methods to aid in the identification of organisms that cannot be cultivated due to unusual growth characteristics or antibiotic treatment or cannot be classified by phenotypic methods (136–141). In one study, results obtained with 16S rRNA gene sequencing and the SmartGene IDNS (Zug, Switzerland) database and software compared favorably to those obtained by conventional phenotypic methods and were better than those obtained with a similar rRNA gene method employing a smaller database for a collection of 300 clinical isolates (137). The performance differences between the two 16S rRNA gene methods highlight the importance of the size and breadth of the database for successful classification. Difficulties in separating E. coli from Shigella with 16S rRNA gene sequencing should be expected. These are genetically the same species and have been maintained as separate taxa for medical expediency. The limited findings reported in the studies above and those reported by others (142) show that a small region of the 16S rRNA gene alone will not provide reliable separation of certain medically relevant members of the Enterobacteriaceae family (E. coli/S. sonnei [137] and Escherichia/Shigella/Hafnia [136]). The incorporation of virulence genes that define the Shigella/ EIEC pathotype should help discriminate it from noninvasive pathotypes or communal E. coli. Another approach that has potential to improve microbial identification involves MS. The Ibis T5000 Biosensor System (Abbott Molecular, Des Plains, IL), which is currently used in nonclinical and research settings, uses multiple regions of the 16S and 23S rRNA genes plus several housekeeping genes to discriminate between species within 6 h (143). Validation studies are needed to assess the performance of this technology on clinical specimens. Simple and cost-effective strategies to use whole-genome sequencing to differentiate bacterial species, determine antibiotic resistance potential, detect virulence determinants, and conduct pathogen surveillance for outbreak detection

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 694

are developing rapidly. The application of this technology in the clinical microbiology setting is reviewed by Didelot et al. (144), who highlight its recent successful use to rapidly characterize the novel E. coli O104:H4 variant that caused the 2011 outbreak of bloody diarrhea in Germany and France (145).

Virulence Testing Extraintestinal E. coli Numerous virulence factors have been identified for extraintestinal E. coli (43), particularly the K1 antigen, but these are usually identified only in epidemiologic studies.

Diarrheagenic E. coli Detection of diarrheagenic pathotypes is typically performed on E. coli colonies chosen from selective or nonselective media. If PCR techniques are used, a sweep of confluent growth from a MAC plate may be screened; if the PCR assay is positive, isolated colonies may then be picked and screened individually. Multiplex PCR assays are capable of simultaneously detecting multiple E. coli pathotypes (146). STEC. Two distinct Shiga toxins, Stx1 and Stx2, also referred to as verocytotoxins 1 and 2, have been described in E. coli. STEC may produce either Stx1 or Stx2 or both toxins. The toxin produced by Shigella dysenteriae serotype 1 was recognized first and is designated Stx, while the toxins in E. coli include an Arabic number in association with “Stx.” The Shiga toxins comprise a family of toxins that have similar biologic, genetic, and structural features. Stx and Stx1 are essentially identical. Subtypes of Stx1 and Stx2 have been identified, and a standardized nomenclature for the toxins and PCR protocols for their detection were recently published by Scheutz et al. (147). The toxin subtypes vary in their association with severe disease, with Stx2a, Stx2c, and Stx2d being associated with bloody diarrhea and HUS (148–150); however, host factors, dose, and other bacterial factors play a role in the severity of disease. The production of Stx or the genes encoding Stx can be detected by a variety of biologic, immunologic, or nucleic acid-based assays (46, 151). FDA-approved diagnostic PCR assays for STEC that discriminate between the two toxin types are now available (see “Nucleic Acid Detection” above). Stx has also been directly detected in the blood of HUS patients by use of flow cytometry, even in the absence of serologic or microbiologic evidence of STEC infection (152). STEC strains represent a spectrum of virulence potentials, ranging from the highly virulent O157:H7 serotype that has been responsible for the majority of outbreak cases to low-virulence serotypes that have been isolated only from nonhuman sources. The presence of additional virulence factors other than Stx correlates with disease potential. The most important of these virulence factors are the intimin adhesin and the type III secretion system encoded by the LEE pathogenicity island (43). The eae gene probe for intimin and the hlyA (E-hlyA or ehxA) gene probe for a plasmid-encoded hemolysin have been the most frequently employed methods to determine virulence potential, but probes for at least 25 different virulence-associated genes have been employed to characterize STEC strains (153, 154). STEC strains have been classified into five “seropathotypes” (A through E) based on the occurrence of serotypes in human disease, in outbreaks, and in severe disease (HUS or hemorrhagic colitis) and on possession of specific virulence genes (155).

37. Escherichia, Shigella, and Salmonella n 695

ETEC. The ST and LT enterotoxins produced by ETEC may be detected by a variety of biologic, immunologic, and nucleic acid-based assays (46). Two distinct ST variants (STh and STp) have been identified in human strains. Strains that produce ST only or ST in combination with LT have caused most ETEC outbreaks in the United States (55). Immunoassays for the identification of ST or LT in culture supernatants of ETEC strains are available from at least two commercial sources (Table 3). The ST EIA assay (Denka Seiken Co., Ltd., Tokyo, Japan; and Oxoid Ltd., Basingstoke, United Kingdom) is a competitive enzyme immunoassay for the detection of ST only (156). A reversed passive latex agglutination assay (VET-RPLA; Oxoid [a similar kit is available from Denka Seiken]) detects both cholera toxin and LT, which are highly related antigenically. The effectiveness of the VET-RPLA may be optimized by use of a culture medium designed for LT production, such as Biken’s medium, rather than the medium recommended by the manufacturer (157). EPEC. EPEC, EAEC, and DAEC can be detected by their characteristic patterns of adherence to HEp-2 or HeLa cells in culture (57). These patterns are also observed on formalin- or glutaraldehyde-fixed cells, obviating the need to prepare cells expressly for the assay (158). EPEC strains are defined on the basis of the A/E histopathology produced on epithelial cells and the lack of Stx (reviewed in references 58 and 43). The A/E phenotype can be detected by tissue culture cell assays or by DNA probe or PCR tests for the eae gene, encoding intimin, or the LEE pathogenicity island. The EAF plasmid of typical EPEC (see above) is detected by use of fragment or oligonucleotide probes or PCR primers (46). Atypical EPEC strains possess only the A/E phenotype (LEE pathogenicity island) but do not possess the EAF plasmid. It is likely that only a subset of atypical EPEC strains comprise true human pathogens, although no tests can reliably identify pathogenic isolates.

late is subcultured. Because of shared invasiveness-related characteristics, these assays also detect Shigella strains. DAEC. DAEC strains were initially defined on the basis of a diffuse adherence pattern to cultured epithelial cells, but this phenotype is not specific for enteric strains (70). Various DNA probes and PCR assays have been proposed for DAEC identification, as reviewed previously (46).

Typing Systems Several methods for subtyping have been used for E. coli O157:H7 isolates. In particular, pulsed-field gel electrophoresis (PFGE) methods and multilocus variable-number tandem-repeat analysis (MLVA) methods are useful (46, 163). A national molecular subtyping network, PulseNet, was established in 1996 by the CDC to facilitate subtyping of bacterial foodborne pathogens, including E. coli O157:H7, Shigella, nontyphoidal Salmonella serotypes, and Listeria monocytogenes (164). Successful detection of outbreaks by this network of state and local public health laboratories is dependent on submission of isolates by clinical laboratories for confirmation and subtyping. Determination of the serotype and the antimicrobial susceptibility pattern is usually adequate for defining outbreak strains of ETEC, EPEC, and EIEC. Plasmid typing or PFGE methods may also be helpful for distinguishing between sporadic isolates and outbreak strains, but neither method has been widely used for these groups of E. coli.

Serologic Tests At present, serodiagnostic tests for diarrheagenic E. coli are valuable only for seroepidemiology surveys and are not useful for the diagnosis of sporadic infections. Assays that measure serum antibody response to LPS have been used to detect STEC infection in culture-negative HUS patients (46). Enzyme-linked immunosorbent assays have been described to detect saliva antibodies to LPS (165) and serum antibodies to the secreted EspB protein in HUS patients (166).

Antimicrobial Susceptibilities EAEC. Several simple assays have been described as surrogates for the cell adherence test for identification of EAEC. These include a simple biofilm formation assay on polystyrene (159) and screening for the presence of a pellicle at the surface of broth media (160). EAEC can be identified more definitively by use of a specific DNA probe (the AA or CVD432 probe) (161), which is superior to tissue culture adherence assays in identifying pathogenic strains of EAEC (65). More recent data suggest that the AA probe corresponds to a putative virulence gene called aatA (162), which is under the control of a regulator termed AggR. AggR, in turn, controls several other virulence factors (66). Thus, the aggR gene (which defines typical EAEC) may represent a superior diagnostic target, although data to support this notion are not available. EIEC. EIEC can be identified by various in vivo assays, immunoassays, and nucleic acid-based assays for invasiveness. EIEC can be detected along with Shigella with two commercial kits, one of which targets the invasion plasmid-associated gene ipaH (See “Nucleic Acid Detection” above). Cell culture invasion assays or DNA-based assays for other invasion-related factors are, for the most part, practical only in research settings (46). Plasmid DNA electrophoresis may be used to detect the large, 120- to 140-MDa plasmid associated with invasiveness, but this plasmid is easily lost when the iso-

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 695

Extraintestinal E. coli

In the past 20 years, E. coli strains producing CTX-M βlactamases and AmpC β-lactamases have emerged as significant causes of extraintestinal infections in the United States and globally (35, 167–170). CTX-M-producing E. coli strains are often isolated from urinary tract infections, both health care and community acquired, and have also been detected in retail meat samples in the United States (35). Recently, a single pandemic clone of multiresistant E. coli ST131 has emerged as the predominant extended-spectrum β-lactamase (ESBL)-positive E. coli strain isolated from urinary tract infections and bacteremia in hospitalized patients and community-acquired infections in the elderly (40, 169– 171). In addition to being resistant to fluoroquinolones, E. coli ST131 is often associated with the CTX-M-15 ESBL (169) and often possesses multiple virulence genes (iha, sat, and iutA). AmpC β-lactamases are problematic for clinical laboratories because these enzymes can interfere with ESBL confirmatory tests, resulting in a false report of cephalosporin susceptibility (169, 170). The AmpC β-lactamase CMY-2 has the broadest geographic distribution and has been found in E. coli and other Enterobacteriaceae (reviewed by Jacoby [172]). Carbapenemases are β-lactamases that confer resistance to the carbapenems, and the Klebsiella pneumoniae carbapen-

696 n BACTERIOLOGY

emase (KPC) is the most frequently encountered enzyme of this class in E. coli and other carbapenem-resistant Enterobacteriaceae. It is important to detect KPC and other carbapenemases in patients colonized with carbapenem-resistant Enterobacteriaceae so that isolation precautions may be instituted to prevent transmission in health care settings (173). Until laboratories can implement the carbapenem interpretive criteria published in the 2014 CLSI guidelines (174), performance of the modified Hodge test (MHT) as described in the guidelines (Table 3C) is advised. If the 2014 CLSI interpretive criteria are used, the MHT does not need to be performed other than for epidemiologic or infection control purposes and no change in the interpretation of carbapenem susceptibility test results is required for MHT-positive isolates. Results of the MHT should be reported to infection control or those requesting epidemiologic information. When identifying carbapenemase-producing isolates for infection control or epidemiologic purposes, a targeted strategy for testing isolates can be used. Selection of isolates for testing in the MHT with the following characteristics is suggested: intermediate or resistant to one or more carbapenems when using the CLSI 2014 interpretive criteria (ertapenem nonsusceptibility is the most sensitive indicator of carbapenemase production) and resistant to one or more agents in cephalosporin subclass III (174). Current automated susceptibility test systems may not be able to accurately detect ESBLs, AmpCs, and KPCs (175, 176). Issues surrounding the testing for these enzymes are reviewed by Thomson (177). In January 2010, the CLSI published interpretative criteria for phenotypically assessing the susceptibility of the Enterobacteriaceae to the cephalosporins and aztreonam (178). Under these guidelines, lower breakpoints were recommended, thereby eliminating the need to perform routine ESBL tests and to edit the results on reports from susceptible to resistant for cephalosporins, aztreonam, or penicillins. No reduction in breakpoints was proposed at that time for cefuroxime (parenteral) and cefepime; however, in 2014, the CLSI substantially revised only the cefepime interpretive criteria (breakpoints) and introduced the “susceptibledose dependent” (SDD) category to replace the “intermediate” category (174). The SDD interpretive category, which has been used successfully to guide antifungal therapy, is a new category for antibacterial susceptibility testing that implies that the susceptibility of an organism is dependent on the dosing regimen that is used in the patient. The CLSI recommends that the following comment be reported with the new cefepime breakpoints: “The interpretive criterion for susceptible is based on a dosage regimen of 1 gram every 12 hours. The interpretive criterion for susceptible-dose dependent is based on dosing regimens that result in higher cefepime exposure, either higher doses or more frequent doses or both, up to approved maximum dosing regimens.” Educating physicians and other stakeholders about the meaning of the SDD category will be important to maximizing the useful life span of cefepime. Additional guidance for implementing the new criteria is discussed in the 2014 CLSI guidelines (174). Until the FDA revises its breakpoints for manufacturers to update their instruments, laboratories using automated antimicrobial susceptibility devices are advised to consult with manufacturers for guidance on how to manually adjust the cefepime breakpoints. A validation study should be performed when altering manufacturers’ settings, and guidelines for performing such a study are described by Clark et al. (179). Implementation of the revised breakpoints and SDD category currently applies only to cefepime susceptibil-

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 696

ity testing of members of the Enterobacteriaceae. The SDD category has been adopted due to the growing need to refine susceptibility reporting to maximize clinicians’ use of available drugs. The CLSI suggests that routine antimicrobial susceptibility testing for Enterobacteriaceae include ampicllin, cefazolin (MIC only), gentamicin, and tobramycin testing. Urinary isolates of E. coli may be tested against fosfomycin and other drugs used only for these infections (trimethoprimsulfamethoxazole). For E. coli and other Enterobacteriaceae isolates recovered from cerebrospinal fluid (CSF), cefotaxime or ceftriaxone should be tested and reported in place of cefazolin. The following antimicrobial agents should not be routinely reported for E. coli or other Enterobacteriaceae isolated from CSF because they are not the drugs of choice and may not be effective for treating CSF infections: agents administered by the oral route only, first- and second-generation cephalosporins (except cefuroxime parenteral), cephamycins, clindamycin, macrolides, tetracyclines, and fluoroquinolones.

Diarrheagenic E. coli STEC Antimicrobial therapy for O157 STEC diarrhea or HUS is controversial: some publications have suggested that antibiotics increase the risk of HUS (48, 180), while a metaanalysis of published reports found no significantly increased risk (181). There is a lack of evidence to support routine antimicrobial susceptibility testing of STEC strains. Until recently, E. coli O157:H7 isolates were almost uniformly susceptible to antimicrobial agents. However, since the early 1990s, O157 and other STEC strains have demonstrated slowly increasing levels of resistance to certain antibiotics, particularly streptomycin, sulfonamides, and tetracycline (http://www.cdc.gov/narms/index.html).

ETEC, EPEC, EAEC, and EIEC Strains and Other Diarrheagenic E. coli Strains Treatment with an appropriate antibiotic can reduce the severity and duration of symptoms of ETEC infection (46). Antimicrobial resistance, particularly to tetracycline, is common among ETEC strains isolated from outbreaks in the United States (55). Antibiotic treatment may be helpful for diarrhea caused by EPEC (46). Most EPEC strains associated with outbreaks are resistant to multiple antimicrobial agents (58). EAEC strains are commonly resistant to most antibiotics, though these strains are typically susceptible to fluoroquinolones. Clinical studies have demonstrated the effectiveness of ciprofloxacin for travelers with diarrhea caused by EAEC (182). Little information about the efficacy of antimicrobial treatment or the prevalence of resistance is available for EIEC or other putative diarrheagenic E. coli strains, but determination of the antimicrobial susceptibility pattern may be helpful in establishing whether the isolates are associated with an outbreak.

Evaluation, Interpretation, and Reporting of Results Extraintestinal E. coli The final written report should include the final Gram stain result, the final identification as E. coli, and the antimicrobial susceptibility test results. Any supplemental tests used to detect resistance markers like KPCs should be highlighted. Laboratories should determine what resistance markers are important for institutional infection control practices.

37. Escherichia, Shigella, and Salmonella n 697

Diarrheagenic E. coli STEC A presumptive diagnosis of an O157 STEC (isolate positive for O157 antigen) or a non-O157 STEC (isolate positive for Shiga toxin) infection should be reported to the clinician as soon as the laboratory obtains this result. The 2009 recommendations for clinical diagnosis of STEC include examples of how to word the final report for results from testing for Shiga toxin immunoassays or PCR for Shiga toxin genes as well as culture results (93; http://www.cdc.gov/mmwr/ preview/mmwrhtml/rr5812a1.htm [Table 2]). When O157 STEC is not found in a specimen, it is advisable to include a comment in reports stating that non-O157 STEC strains can cause diarrhea and HUS. Cases of STEC infection and HUS should be reported to public health authorities. O157 STEC isolates or specimens in which Shiga toxin or STEC genes are detected (e.g., stool, other primary specimens, and enrichment broths) should be forwarded to a local or state public health laboratory (93).

ETEC, EPEC, EAEC, and EIEC Strains Generally, the ETEC, EPEC, EAEC, and EIEC classes of diarrheagenic E. coli are identified only during outbreak investigations. A laboratory reporting these results, which usually will be a retrospective diagnosis obtained by a reference laboratory, should provide an explanation of the clinical significance of these organisms and may refer the clinician to the reference laboratory for further information. All suspected outbreaks should be reported to public health authorities.

SHIGELLA Taxonomy Shigella is classified in the family Enterobacteriaceae, which is addressed in chapter 38 of this Manual (2, 3). There are four subgroups of Shigella that historically have been treated as species: subgroup A as S. dysenteriae, subgroup B as Shigella flexneri, subgroup C as S. boydii, and subgroup D as Shigella sonnei. From a genetic standpoint, the four species of Shigella, with the exception of S. boydii 13, and E. coli represent a single genomospecies (183–185). Using a genetic definition for species, the four species of Shigella would be regarded as serologically defined anaerogenic biotypes of E. coli. The current nomenclature for Shigella organisms is maintained largely for medical purposes because of the useful association of the genus epithet with the distinctive disease (shigellosis) caused by these organisms. The G+C content of the DNA is 49 to 53 mol%, and the type species for the genus is S. dysenteriae (Shiga 1898) (5). S. boydii 13 strains were first described in 1952 and then added to the Shigella scheme in 1958 (186). Early findings

from DNA-DNA hybridization showed that these strains represent a new species (185); however, it was not until recently that findings from phylogenetic studies showed that they cluster in a neighbor-joining tree with E. albertii, a newly described species of Escherichia associated with diarrheal disease in Bangladeshi children (8, 30).

Description of the Genus The genus Shigella is composed of nonmotile bacteria that conform to the definition of the family Enterobacteriaceae (2, 187). Species in this genus are Gram-negative rods that grow well on MAC. All strains of Shigella spp. are nonmotile; do not decarboxylate lysine; do not utilize citrate, malonate, or sodium acetate (with exceptions for S. flexneri); and do not grow in KCN or produce H2S. Compared with Escherichia, Shigella strains are less active in their use of carbohydrates (Table 4). All ferment D-glucose without the production of gas (a few exceptions produce gas, e.g., certain strains of S. flexneri serotype 6 and S. boydii serotype 14). S. sonnei strains ferment lactose and sucrose on extended incubation, but other species generally do not use these substrates in conventional medium. Salicin, adonitol, and myo-inositol are not fermented. There are numerous identical and reciprocal serologic reactions between Shigella and E. coli (188).

Epidemiology and Transmission Humans and other large primates are the only natural reservoirs of Shigella bacteria. Most transmission is by person-toperson spread, but infection is also caused by ingestion of contaminated food or water. Shigellosis is most common in situations in which hygiene is compromised (e.g., child care centers and other institutional settings). In developing populations without running water and indoor plumbing, shigellosis can become endemic. Sexual transmission of Shigella among men who have sex with men also occurs. In the United States, an estimated 495,000 cases of shigellosis occur each year, with 38 deaths (47). Up to 15% of all U.S. cases of shigellosis are related to international travel. Most infections in the United States and other developed countries are caused by S. sonnei; S. flexneri is the secondmost-common subgroup (http://www.cdc.gov/nationalsur veillance/shigella-surveillance.html). In the developing world, the majority of endemic dysentery cases are caused by S. flexneri, with the balance of cases caused by subgroups that vary temporally and geographically (189–191). Epidemic dysentery is most commonly caused by S. dysenteriae 1, whose prevalence rises dramatically during outbreak periods and then falls as the epidemic resolves. Infection with S. dysenteriae 1 is associated with high rates of morbidity and mortality in developing countries, particularly when antimicrobial resistance or misdiagnosis delays appropriate treatment. In the United States and other developed countries, S. sonnei is endemic and causes large, protracted outbreaks in day care

TABLE 4 Differentiation of E. coli and Shigella Test Resulta of test with:

Shigella Inactive E. colib E. coli

Lysine decarboxylase − d +

Motility

Gas from glucose

Acetate utilization

− − +

− − +

− d +

Mucate Lactose − d +

− d +

a Abbreviations and symbols: +, 90% or more positive within 2 days; −, no reaction (90% or more) in 7 days; d, different reactions [+, (+), +]. Adapted from Ewing (2). b Nonmotile, anaerogenic biotypes sometimes referred to as Alkalescens-Dispar bioserotypes.

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 697

698 n BACTERIOLOGY

centers (192–194) and among men who have sex with men (195, 196). The protracted nature of these outbreaks is attributed to a large number of asymptomatically infected individuals in the population and the tendency for secondary spread (197). For most individuals, antibiotic treatment reduces the number of symptomatic days and the length of shedding (198); however, the decision to treat is nuanced and is influenced by a variety of factors, including severity of disease, immune status of the host, likelihood of transmission to others, preservation of the microbiome, and development of antibiotic resistance. Because resistance to ampicillin and trimethoprim-sulfamethoxazole is common in day care-associated S. sonnei infections, azithromycin has been recommended for treatment of pediatric shigellosis. The effectiveness of this recommendation will require careful monitoring in light of an outbreak in 2012 caused by a S. sonnei strain with reduced susceptibility to azithromycin (199).

Clinical Significance Members of the genus Shigella have been recognized since the late 19th century as causative agents of bacillary dysentery. Shigella causes bloody diarrhea (dysentery) and nonbloody diarrhea. Shigellosis often begins with watery diarrhea accompanied by fever and abdominal cramps but may progress to classic dysentery with scant stools containing blood, mucus, and pus. Ulcerations, which are restricted to the large intestine and rectum, typically do not penetrate beyond the lamina propria. Bloodstream infections can occur but are rare. Appropriate antimicrobial therapy will decrease the duration, transmission, and severity of symptoms and is customarily prescribed based on the severity of illness or the need to protect close contacts. Patients in certain occupations (i.e., food handlers, child care providers, and health care workers) and children who attend day care often are required to have a documented negative stool culture following treatment. The infectious dose is low (1 to 100 organisms), and the incubation period is 1 to 4 days. Shigellae are shed in stools for several days to several weeks after illness, and persons who receive appropriate antimicrobial therapy will be culture negative at 72 h (198). All four subgroups of Shigella are capable of causing dysentery, but S. dysenteriae serotype 1 has been associated with a particularly severe form of illness thought to be related in part to its production of Shiga toxin. Infection can occasionally be asymptomatic, particularly infection with S. sonnei strains. Complications of shigellosis include HUS, which is associated with S. dysenteriae 1 infection, and reactive arthritis or Reiter’s chronic arthritis syndrome, which is associated with S. flexneri infection (200). The identification of Shigella species is important for both clinical and epidemiologic purposes.

Collection, Transport, and Storage of Specimens See “Collection, Transport, and Storage of Specimens” in the Escherichia section.

not practical and has not been validated, and no commercial FDA-approved kits are available.

Nucleic Acid Detection There are four FDA-cleared nucleic acid detection methods for clinical diagnosis of Shigella infections. See “Nucleic Acid Detection” under “Direct Examination” in the Escherichia section for more information.

Isolation Procedures Enrichment and Plating Media There is no reliable enrichment medium for all Shigella isolates, but Gram-negative broth and Selenite broth (SEL) are frequently used. For the optimal isolation of Shigella, two different selective media should be used: a general purpose plating medium of low selectivity (e.g., MAC) and a more selective agar medium (e.g., xyloselysine-deoxycholate agar [XLD]). Statens Serum Institut enteric medium, deoxycholate-citrate agar, and Hektoen enteric agar (HE) are suitable alternatives to XLD as media with moderate to high selectivities. Salmonellashigella agar should be used with caution because it inhibits the growth of some strains of S. dysenteriae 1.

Screening Procedures Shigella strains appear as lactose- or xylose-nonfermenting colonies on the isolation media described above. S. dysenteriae 1 colonies may be smaller on all of these media, and these strains generally grow best on media with low selectivities (e.g., MAC). S. dysenteriae 1 colonies on XLD agar are frequently very tiny, unlike those of other Shigella species. S. sonnei colonies often appear flattened and spread out on blood agar plates. Suspect colonies may be screened phenotypically on Kligler iron agar (KIA) or triple sugar iron agar (TSI). Shigella species characteristically produce an alkaline slant because strains do not ferment lactose (or sucrose) and do not produce gas or H2S. A few strains of S. flexneri 6 and a very few strains of S. boydii produce gas in KIA or TSI. The motility and lysine decarboxylase tests are characteristically negative for Shigella and can be used to further screen isolates before serologic testing (Table 4). Serologic testing should be performed as recommended by the manufacturer of the product being used. Due to the many antigenic cross-reactions between E. coli and Shigella, Shigella antisera are typically validated to be used in the identification of bacteria that biochemically conform to the definition of Shigella. For this reason, the initial screening of isolates for which there is no phenotypic evidence that they conform to the definition of Shigella is discouraged. Isolates that react appropriately with the screening tests should then be identified with a complete set of phenotypic tests, with automated systems or self-contained commercial kits being satisfactory, and should be tested with grouping antisera. Confirmation requires both phenotypic and serologic identification, and laboratories that do not perform both types of tests should send Shigella isolates to a reference laboratory for confirmation.

Direct Examination Identification Microscopy Shigella cannot readily be distinguished from other Gramnegative rods by staining or microscopy methods.

Antigen Detection Because there is no single somatic antigen common to all Shigella strains, antigen detection in clinical specimens is

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 698

Phenotypic Identification Shigellae are biopathotypes of E. coli that are identified conventionally using a combination of biochemical tests and O serogrouping (188). Shigella and inactive E. coli (anaerogenic or lactose-nonfermenting) strains are frequently difficult to distinguish by routine phenotypic tests.

37. Escherichia, Shigella, and Salmonella n 699

See Table 4 for the phenotypic reactions characteristic of Shigella spp. Although S. dysenteriae and S. sonnei are phenotypically distinct, S. flexneri and S. boydii are often phenotypically indistinguishable, so serologic grouping is essential to identify them to the species (subgroup) level. The ability of commercial methods to identify Shigella was assessed several years ago by O’Hara and Miller (122– 124), who found that these systems correctly identified only 50 to 70% of the 10 Shigella strains in their test panel and misidentified several other members of the Enterobacteriaceae as Shigella. MALDI-TOF MS systems for the rapid identification of bacteria were similarly disappointing and could not correctly identify Shigella (they identified Shigella as E. coli) (126, 127). This is not surprising because the accurate identification of Shigella requires performing serogrouping in Shigella-specific antisera in addition to biochemical profiling. Molecular assays targeting Shigella virulence genes (e.g., ipaH, ipaB, ipaC, inv, and ial) have been very useful for detecting Shigella/EIEC and identifying potential new serotypes.

Serotyping Serotyping is essential for the identification of Shigella. Three of the four subgroups, A (S. dysenteriae), B (S. flexneri), and C (S. boydii), are made up of a number of serotypes. Subgroup A has 15 serotypes; subgroup B has 8 serotypes (with serotypes 1 to 5 subdivided into 11 subserotypes); and subgroup C has 19 serotypes, numbered 1 through 20, with S. boydii 13 reclassified as E. albertii. Subgroup D (S. sonnei) is made up of a single serotype. Subgroups A and C are rare. Several provisional Shigella serotypes have also been described, which are held sub judice until findings from the characterization of representative isolates show them to be unique and of sufficient prevalence to merit inclusion in the Shigella scheme. Antisera for the identification of provisional serotypes are typically available only at reference laboratories. Serotyping is typically performed by slide agglutination with polyvalent somatic (O) antigen grouping sera, followed, in some cases, by testing with monovalent antisera for specific serotype identification. Monovalent antiserum to S. dysenteriae 1 is required to identify this serotype and is not widely available. Because of the potentially serious nature of illness associated with this serotype, isolates that agglutinate in subgroup A reagent should be sent to a reference laboratory immediately for further serotyping. Phenotypically typical Shigella isolates that agglutinate poorly or that do not agglutinate at all should be suspended in saline and heated in a water bath at 100°C for 15 to 30 min. After cooling, the antigen suspension should be tested in normal saline to determine if it is rough (agglutinates spontaneously). If the heated and cooled suspension is not rough, it may then be retested for agglutination in antisera.

Typing Systems A variety of methods have been used to subtype Shigella, including colicin typing (particularly for S. sonnei), plasmid profiling, restriction fragment length polymorphism analysis, PFGE, and ribotyping (201). More recently, MLVA and a single nucleotide polymorphism typing scheme for the rapid and discriminatory typing of Shigella have been developed (202, 203). For an overview of the epidemiologic use of typing methods, see chapter 10.

Serologic Tests Several serodiagnostic assays based on different antigens possessed by Shigella have been described (204, 205). These

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 699

assays are practical only in research settings for seroepidemiology surveys and are not currently used for the diagnosis of infection in individual patients.

Antimicrobial Susceptibilities Because of the widespread antimicrobial resistance among Shigella strains, all isolates should undergo susceptibility testing. The CLSI recommends that susceptibility results be reported routinely for only ampicillin, trimethoprim-sulfamethoxazole, and a fluoroquinolone and warns that Shigella should not be reported as susceptible to first- and secondgeneration cephalosporins, cephamycins, and aminoglycosides, because these drugs are often not effective clinically (206). Because of widespread resistance in the United States, ampicillin and trimethoprim-sulfamethoxazole, two safe drugs that were the most commonly prescribed for treating children with S. sonnei infections, are no longer options for empiric treatment. Macrolides, in particular azithromycin, are being used to treat these infections, but there are no interpretive criteria for antimicrobial susceptibility testing for Shigella, making it problematic to monitor for development of resistance (199). Reporting of susceptibility results to the clinician is particularly important for S. dysenteriae 1 isolates. Infections caused by these strains are often acquired during international travel to areas where most strains are multidrug resistant (207). In many areas of Africa and Asia, S. dysenteriae 1 strains are resistant to all locally available antimicrobial agents, including nalidixic acid (190, 208), and fluoroquinolone-resistant strains have been reported in Asia (191, 209, 210).

Evaluation, Interpretation, and Reporting of Results A preliminary report of suspected Shigella infection may be issued if serologic or biochemical screening tests are positive or consistent with Shigella. If serotyping results (O antigen determination) are available, these should also be reported, particularly if the isolate is S. dysenteriae 1. All Shigella isolates should be tested for antimicrobial susceptibility. Before issuing a final report, isolates should be confirmed by both biochemical and serologic methods, because the definition of Shigella is dependent on both the biochemical profile and O antigen expressed. Isolates, particularly those from individuals with dysentery-like illness, that are biochemically identified as Shigella but that are serologically negative may be new serotypes of Shigella and should be sent to a reference laboratory for further characterization. Isolates from sites other than the gastrointestinal tract that resemble Shigella should be scrutinized carefully for gas production and other differentiating characteristics because extraintestinal Shigella infections are rare. These isolates should be sent to a reference laboratory for confirmation because they are more likely to be anaerogenic E. coli, certain strains of which may cross-react with Shigella antiserum.

SALMONELLA Taxonomy Members of the genus Salmonella are classified in the family Enterobacteriaceae (2, 3). Species of this genus are motile, Gram-negative, facultative rods. Salmonellae are typically defined by their ability to use citrate as a sole carbon source and lysine as a nitrogen source and by the production of H2S on triple sugar agar; exceptions to these traits are used to define species, subspecies, and some serotypes (2, 211).

700 n BACTERIOLOGY

The genus Salmonella is composed of two species, Salmonella enterica and Salmonella bongori (212, 213). S. enterica is subdivided into six subspecies: S. enterica subsp. enterica, often called subspecies I; S. enterica subsp. salamae, or subspecies II; S. enterica subsp. arizonae, or subspecies IIIa; S. enterica subsp. diarizonae, or subspecies IIIb; S. enterica subsp. houtenae, or subspecies IV; and S. enterica subsp. indica, or subspecies VI. The type species is S. enterica subsp. enterica. Subspecies IIIa and IIIb represent organisms originally described in the genus “Arizona”; subspecies IIIa contains the monophasic strains and subspecies IIIb the diphasic strains of “Arizona” (214). Despite their common history, subspecies IIIa and IIIb are more closely related to some of the other subspecies of S. enterica than they are to each other and thus should be considered separate entities (215, 216). Genome analysis of the salmonellae has revealed a high degree of genetic variability. Serotypes within S. enterica subspecies I have been shown to differ by as much as 10% in gene content (i.e., presence or absence of whole genes) (217, 218). Recombination, particularly among strains of S. enterica subspecies I, likely contributes to this diversity (219). The use of whole-genome sequence analysis to characterize strains continues to expand, with >500 complete or partial sequences from Salmonella strains available as of the end of 2013 (http://www.ncbi.nlm.nih.gov/genome/ browse/). Characterization of whole-genome sequences continues to increase our understanding of Salmonella biology and phylogeny. Analysis of genome sequences revealed two distinct lineages among S. enterica subsp. enterica; differences in gene content were noted that may contribute to differ-

ences in ecology or transmission (220, 221). The development of tools for the analysis of whole-genome sequences based on single nucleotide polymorphisms, core genes, or other markers has already begun and will undoubtedly lead to a revolution in epidemiologic strain typing (221–223).

Description of the Genus Subspecies I strains are commonly isolated from humans and warm-blooded animals. Subspecies II, IIIa, IIIb, IV, and VI strains and S. bongori are usually isolated from coldblooded animals and the environment. Non-subspecies I strains are typically considered rare human pathogens; they make up about 1 to 2% of Salmonella isolates reported to the U.S. National Salmonella Surveillance System (http:// www.cdc.gov/nationalsurveillance/salmonellasurveillance.html). The phenotypic traits useful for identification of Salmonella are given in Table 5 and for subspecies differentiation are given in Table 6. The nomenclature employed to describe the genus Salmonella was problematic for many years due to the use of multiple schemes in the literature and the historical practice of considering different serotypes of Salmonella to be different species. The publication of Judicial Opinion 80 in the International Journal of Systematic and Evolutionary Microbiology in 2005 (212) hopefully served to clarify nomenclatural issues regarding the genus Salmonella, and the conventions set forth in that opinion are used here. Salmonella history and nomenclature are reviewed at http://www.bacterio.net/ salmonella.html.

TABLE 5 Phenotypic traits useful for differentiating Salmonella from other Enterobacteriaceae and identifying Salmonella serotypes Typhi and Paratyphi Aa Reaction for: Test

TSI H2S (TSI) Indole Methyl red Voges-Proskauer Citrate (Simmons) Urea Lysine decarboxylase Arginine dihydrolase Ornithine decarboxylase Motility Mucate Malonate L(+)-Tartrate (d-tartrateb) Growth in KCN Glucose Lactose Sucrose Salicin Dulcitol Inositol Sorbitol o-Nitrophenyl-β-D-galactopyranoside Galacturonate

Nontyphoidal Salmonella subsp. I

Salmonella serotype Typhi

Salmonella serotype Paratyphi A

K/A, g + − + − + − + + + + + − + − A, g − − − A, g d A, g − −

K/A +weak − + − − − + d − + − − + − A − − − − − A − −

K/A, g − or +weak − + − − − − (+) + + − − − − A, g − − − A, g2 days − A, g − −

a Reactions after incubation at 37°C. Abbreviations and symbols: +, 90% (or more) of strains were positive within 2 days; (+), positive reaction after 3 or more days; −, no reaction (90% or more) in 7 days; A, acid; d, different reactions [+, (+), −]; g, gas; K, alkaline slant. Adapted with permission of the publisher from reference 2. b Sodium potassium tartrate (2).

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 700

37. Escherichia, Shigella, and Salmonella TABLE 6

n

701

Phenotypic traits useful for differentiating Salmonella species and subspeciesa

Reaction for: Species or subspecies (no. of strains tested) Dulcitol Lactose ONPG Salicin Sorbitol Galacturonate Malonate Mucate S. enterica I (650) S. enterica II (146) S. enterica IIIa (120) S. enterica IIIb (155) S. enterica IV (120) S. enterica VI (9) S. bongori (16)

Growth in KCN

Gelatin (strip)

L(+)-Tartrate

(d-tartratej)

+

−

−

−

+

−

−

+

−

−

+

+

−

−f

−

+

+

+

+

−

+

−

−

−c

−

−

+

−

+

+

−

+

−

−

−d

−

−

+

+

+

−i

−

+

−

−

−

−

+h

+

+

−

−

+

+

−

db

de

dg

−

−

+

−

+

−

+

−

+

−

+

−

+

+

−

+

+

−

−

a Reactions after incubation at 37°C. Abbreviations and symbols: +, 90% (or more) of strains were positive within 1 or 2 days; (+), positive reaction after 3 or more days; −, no reaction (90% or more) in 7 days; d, different reactions [+, (+), −]; ONPG, o-nitrophenyl-β-D-galactopyranoside. Adapted with permission of the publisher from reference 2. b A total of 67% were positive. c A total of 15% were positive. d A total of 85% were positive. e A total of 22% were positive. f A total of 15% were positive. g A total of 44% were positive. h A total of 60% were positive. i A total of 30% were positive. j Sodium potassium tartrate (2).

Epidemiology and Transmission Salmonella organisms are isolated most frequently from the intestines of humans and animals. Some serotypes are isolated only from humans (e.g., Salmonella serotype Typhi), while others (e.g., Salmonella serotype Gallinarum and Salmonella serotype IV 48:g,z51:− [formerly serotype Marina]) are strongly associated with certain animal hosts. Members of this genus can be isolated from feces-contaminated food or water but probably do not occur as free-living organisms in the environment. Historically, Salmonella has been considered a pathogen of meat and poultry products, but it has recently been associated with other food vehicles, such as fresh produce and manufactured products (224– 226).

Salmonella Serotypes Salmonella serotyping is a subtyping method based on the immunologic characterization of three surface structures: O antigen, which is the outermost portion of the LPS layer that covers the bacterial cell; H antigen, which is the filament portion of the bacterial flagella; and Vi antigen, which is a capsular polysaccharide present in specific serotypes. Serotyping of Salmonella is commonly performed to facilitate public health surveillance for Salmonella infections and to aid in the recognition of outbreaks. While molecular methods such as PFGE and MLVA have become the cornerstone of public health subtyping (227, 228), serotyping remains an important tool. The serotype of an isolate often correlates with a particular disease syndrome or food vehicle, making serotype data particularly useful in identifying cases and defining outbreaks. For example, Salmonella serotype Typhi causes typhoid fever, a more severe disease syndrome than those caused by most other Salmonella serotypes. Salmonella serotype Enteritidis is often associated with infections acquired from chicken or egg products (223). Furthermore, Salmonella serotyping is

MCM 11th Edition X : MCM11$CH37 03-23-15 12:29:08

Page 701

performed worldwide and has aided in the recognition of international outbreaks (229).

Clinical Significance Strains of Salmonella are categorized as typhoidal and nontyphoidal, corresponding to the disease syndrome with which they are associated. Strains of nontyphoidal Salmonella usually cause intestinal infections (accompanied by diarrhea, fever, and abdominal cramps) that often last 1 week or longer (230). Less commonly, nontyphoidal Salmonella can cause extraintestinal infections (e.g., bacteremia, urinary tract infection, or osteomyelitis), especially in immunocompromised persons. Persons of all ages are affected, but the incidence is highest in infants and young children. Salmonella is ubiquitous in animal populations, and human illness is usually linked to foods. Salmonellosis is also transmitted by direct contact with animals, by water, and occasionally by human contact. Each year, an estimated 1 million cases of illness and 378 deaths are caused by nontyphoidal salmonellosis in the United States (47). Typhoid fever, caused by Salmonella serotype Typhi, is a serious bloodstream infection common in the developing world. However, it is rare in the United States, where on average an estimated 1,800 cases, with 60% of U.S. cases are related to foreign travel (47). Typhoid fever typically presents with a sustained debilitating high fever and headache. Adults characteristically present without diarrhea. Illness is milder in young children, where it may manifest as nonspecific fever. Humans are the only reservoir for Salmonella serotype Typhi, indicating that this serotype is adapted to the human host; healthy carriers have been noted. Typhoid fever typically has a low infectious dose (10%; when patients with typhoid fever are treated with appropriate antibiotics, the

37. Escherichia, Shigella, and Salmonella

rate should be 2 months, occurs in only 10% of cases. Overall, 50% of cases are bacteremic; the presence of >100 CFU/ml of blood and a blood culture showing growth in the first 24 h of incubation are markers for high mortality. At the mild end of the clinical spectrum of melioidosis is presentation with a skin ulcer or abscess without systemic illness (99). Other common presentations with or without bacteremia are genitourinary infections, septic arthritis, and osteomyelitis (83, 84, 96). Prostatic abscesses are especially common (100). Abscesses can also occur in spleen, liver, kidneys, and adrenal glands. Parotitis, lymphadenitis, sinusitis, orchitis, myositis (especially psoas abscess), mycotic aneurysms, and pericardial and mediastinal collections have all been described. Lesions can be frankly purulent and may include microabscesses or granulomas or a combination of these features. Clinical meningitis is rare, but melioidosis encephalomyelitis syndrome (96) and brain abscesses have also been reported. The one presentation that has yet to be described is B. pseudomallei endocarditis. B. mallei is the etiologic agent of glanders, a highly communicable disease of livestock, particularly horses, mules, and donkeys. It can be transmitted to humans and is also identified as a potential agent of bioterrorism. Unlike B. pseudomallei, B. mallei is a host-adapted pathogen that does not persist in the environment outside its host. Glanders has been eradicated from most countries, but enzootic foci persist in the Middle East, Asia, Africa, and South and Central America. The only human case of glanders in the past 50 years in the United States was a recent laboratoryacquired case in a biodefense scientist (101). Like melioidosis, human glanders can be acute or chronic, with the clinical presentation and course depending on the mode of infection, the inoculation dose, and host risk factors. Respiratory inoculation can result in pneumonia with potential for dissemination to internal organs and septicemia. Cutaneous inoculation can result in skin nodules and regional lymphadenitis, also with potential for disseminated disease. Involvement of nodes, both mediastinal and peripheral, is much more common in glanders than in melioidosis, often with suppurative abscesses in untreated cases.

S. maltophilia S. maltophilia, although typically not pathogenic for healthy persons, is a well-known opportunistic human pathogen. It is among the most common causes of wound infection due to trauma involving agricultural machinery (102). It is also an important nosocomial pathogen associated with substantial morbidity and mortality, particularly in debilitated or immunocompromised patients and patients requiring ventilatory support in intensive care units (103). The incidence of human infection appears to have increased in recent years, and a variety of clinical syndromes have been described, including bacteremia, pneumonia, urinary tract infection, ocular infection, endocarditis, meningitis, soft tissue and wound infection, mastoiditis, epididymitis, cholangitis, osteochondritis, bursitis, and peritonitis (104, 105). Septicemia can be accompanied by ecthyma gangrenosa, a skin lesion more commonly associated with P. aeruginosa and Vibrio spp. The incidence of S. maltophilia respiratory tract infection in persons with CF also appears to be increasing (106); however, the unreliability of historical data limits firm conclusions. Approximately 14% of CF patients included in the

43. Miscellaneous Gram-Negative Bacteria n 795

Cystic Fibrosis Foundation’s patient registry were reported to be culture positive for S. maltophilia in 2011 (70). In large, multicenter clinical trials, however, S. maltophilia was found in a larger proportion of CF patients, being second only to P. aeruginosa in frequency of isolation from study subjects. Infection or colonization was most frequently transient, with 30% of subjects having at least one sputum culture positive for S. maltophilia during the course of 6 months (107). Several case-control studies have drawn conflicting conclusions regarding the role that S. maltophilia plays in contributing to pulmonary decline in CF (108).

Ralstonia and Cupriavidus spp. As described above, several former Ralstonia species are now assigned to the genus Cupriavidus (5). Among the species in these two genera, R. pickettii is best known with respect to human infection. Older reports describe this species as being recovered from a variety of clinical specimens and as causing various infections including bacteremia, meningitis, endocarditis, and osteomyelitis (109). R. pickettii also has been identified in pseudobacteremias and nosocomial outbreaks due to contamination of intravenous medications, “sterile” water, saline, chlorhexidine solutions, respiratory therapy solutions, and intravenous catheters (110, 111). This species has also been recovered from the respiratory tract of persons with CF. However, R. pickettii is easily confused with Pseudomonas fluorescens and members of the B. cepacia complex based on phenotype (17, 112). Furthermore, several R. pickettii-like species are also now known to be involved in human infection, particularly in CF (113). Thus, the role of R. pickettii as a human pathogen is difficult to assess based on historical data. R. mannitolilytica (formerly known as R. pickettii biovar 3/ ‘thomasii’) was recently described as causing nosocomial outbreaks and a case of recurrent meningitis (17). This species accounts for the majority of Ralstonia infections in CF patients, being found in more than twice as many CF patients as those infected with R. pickettii (113). R. insidiosa and C. respiraculi are species that are also recovered from persons with CF (18, 21). C. gilardii has been recovered from cerebrospinal fluid (20), and cases of C. pauculus bacteremia, peritonitis, and tenosynovitis have been reported (19). Both of these species may be found in sputa from patients with CF (113). Although Cupriavidus metallidurans and Cupriavidus basilensis are not known to cause other human infection, they too have been recovered recently from sputum cultures from patients with CF (113). Despite these observations, the roles of Ralstonia and Cupriavidus species in human infection, particularly in persons with CF, require further elucidation.

Other Genera In general, Brevundimonas, Comamonas, Delftia, Acidovorax, and Pandoraea spp. infrequently cause human infection. Interest in these species focuses primarily on their roles as plant pathogens or in studies of microbial biodiversity and biodegradation. Brevundimonas spp. are occasionally recovered from clinical specimens (114). B. vesicularis bacteremia in patients with various underlying illnesses has been reported (115), and the organism has been recognized in cervical specimens because of its ability to produce bright orange colonies on ThayerMartin agar. B. diminuta has been recovered from blood, urine, and pleural fluid from patients with cancer (116). Among the Comamonas species, C. testosteroni has been implicated most often in human infection, with reports describing endocarditis, meningitis, and catheter-associated bacteremia due to this species (117–119). D. acidovorans has

similarly been reported to cause infection, being identified in cases of bacteremia, endocarditis, ocular infection, and suppurative otitis (120). Acidovorax spp. have been isolated from a variety of clinical sources, including blood from a patient with hematological malignancy (121). Acidovorax spp., C. testosteroni, and D. acidovorans have also been recovered from sputa of persons with CF (122; J. J. LiPuma, unpublished data); however, the roles of these species in contributing to lung disease in CF have not been established. In addition to causing infection in CF patients (123, 124), Pandoraea spp. have been recovered from blood and from patients with chronic obstructive pulmonary disease or CGD (125). Although the roles of these species in contributing to poor outcomes in persons with underlying diseases are unclear, a recent report describes sepsis, multiple organ failure, and death in a patient who underwent lung transplantation for sarcoidosis (126).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS The genera described in this chapter include organisms that can survive in a variety of hostile environments and at temperatures found in clinical settings. Therefore, standard collection, transport, and storage techniques as outlined in chapter 18 are sufficient to ensure the recovery of these organisms from clinical specimens. Recovery of B. pseudomallei for the diagnosis of melioidosis is increased by the additional collection of throat and rectal swabs into selective media (see “Isolation Procedures” below) and by collecting larger-than-standard volumes of cerebrospinal fluid for culture in suspected neurological melioidosis (127).

DIRECT EXAMINATION Members of these genera have similar morphologies and, with the exception of B. pseudomallei, are not easily distinguished from each other on the basis of Gram staining. B. pseudomallei may appear as small, Gram-negative bacilli with bipolar staining, making the cells resemble safety pins (Fig. 1). This may increase the index of suspicion for the presence of B. pseudomallei, but the sensitivity and specificity of this appearance are not high enough to be relied on for a presumptive clinical diagnosis. Although PCR-based assays have been described for the identification of B. cepacia complex species, B. pseudomallei, B. gladioli, several Ralstonia and Cupriavidus species, Pandoraea species, and S. maltophilia following culture and isolation (see “Identification” below), the use of PCR for direct detection of these species in clinical specimens remains a research tool (128, 129). Studies of CF sputum samples have indicated that some specimens may be PCR positive but culture negative for certain B. cepacia complex species, raising important questions about the natural history of infection in CF. However, the sensitivities and specificities of such PCR assays for the intended target species are difficult to determine in the absence of reliable gold standards. Assays employing real-time PCR or culture-independent strategies employing next-generation bacterial genome sequencing are likely to provide alternative approaches to direct detection of these species in clinical specimens in the near future. Because septicemia with B. pseudomallei is frequently fatal and death often occurs in the first few days after presentation to hospital prior to the availability of culture results, several rapid direct-detection methods have been developed, including urinary antigen detection using latex agglutination and enzyme immunoassay as well as direct fluorescent-antibody

796 n

BACTERIOLOGY

have high sensitivities and specificities with pure bacterial cultures (135–137). Two clinical evaluations of real-time PCR have met with mixed results, with a sensitivity of 91% in one study using an assay targeting a gene in the TSS1 cluster (129) and a considerably lower sensitivity of 61% in a second study that used an assay targeting the 16S rRNA gene (138). Sensitivity of PCR is highest for pus and other body fluids but is low for blood, most likely reflecting a bacterial copy number effect. Various methods have been assessed for optimal DNA extraction from clinical samples (139), and comparisons have been made between the published real-time PCR targets, with the TTS1-orf2 assay found to be superior in detecting B. pseudomallei directly from clinical specimens (140). Loop-mediated isothermal amplification is easy and quick to perform and needs minimal equipment, with amplification being achieved at a fixed temperature in a water bath or heating block. This procedure has been developed and evaluated for the detection of B. pseudomallei and diagnosis of melioidosis (141). The assay was sensitive and specific for the laboratory detection of B. pseudomallei and had 100% specificity when applied to clinical samples but had a very low diagnostic sensitivity (44%). At present, molecular assays are not sufficiently sensitive to replace conventional culture.

ISOLATION PROCEDURES

FIGURE 1 (a) Gram stain of B. pseudomallei in a blood culture; (b) Gram stain of B. pseudomallei from a colony on blood agar. doi:10.1128/9781555817381.ch43.f1

(DFA) staining (130–132). Antibodies raised against heatkilled whole cells of B. pseudomallei have been used to prepare a reagent for DFA staining. When this DFA reagent was used to stain clinical specimens from patients with suspected melioidosis, using a rapid assay that took 10 min to prepare and read, it showed a sensitivity of 66% and specificity of 99.5% (132). A DFA assay has also been described based on a monoclonal antibody specific to B. pseudomallei and Alexa Fluor 488 (133). The reagents described in the literature are largely prepared in-house and are not available commercially. Evaluation of a conventional PCR assay targeting the 16S rRNA gene for the detection of B. pseudomallei in clinical specimens indicated that the assay was sensitive but lacked specificity, resulting in positive predictive values of only 70% (134). Real-time PCR assays targeting B. pseudomallei genes encoding 16S rRNA, flagellin (fliC), ribosomal protein subunit S21 (rpsU), or type III secretion system (TTS) genes have been developed and been shown to

The species discussed in this chapter grow well on standard laboratory media such as 5% sheep blood and chocolate agars. Such media can be used to recover the organisms from sterile fluid or tissue where a mixed biota is not anticipated (see chapter 18). All species that have been reported to be recovered from blood, including B. pseudomallei, grow in brothbased blood culture systems within the standard 5-day incubation period, so that special blood culture techniques such as lysis-centrifugation and extended incubation periods are not required. The use of selective media facilitates the isolation of these organisms from specimens with mixed microbiota. With the exception of B. vesicularis, MacConkey agar can be used to isolate most species of these genera. Burkholderia species grow on MacConkey agar (Fig. 2a), but the use of specific selective media with the ability to inhibit P. aeruginosa is preferred for the isolation of B. cepacia complex and B. pseudomallei. Several selective media have been described, and some are commercially available. A multicenter comparison of three media, PC (for Pseudomonas cepacia) agar (BD Diagnostics, Franklin Lakes, NJ) (142), OFPBL (for oxidation-fermentation base–polymyxin B–bacitracin–lactose) agar (BD Diagnostics) (143), and BCSA (for B. cepacia selective agar; Hardy Diagnostics, Santa Maria, CA) (25), showed that BCSA was superior, being both more sensitive (more B. cepacia isolates were recovered) and more specific (fewer other types of organisms grew) than PC or OFPBL agar (25, 144). The sensitivities of TB-T (for trypan blue-tetracycline), PC-AT (for Pseudomonas cepacia azelaic acid), and BCSA (25, 144) were also compared with those of three commercial media, i.e., B. cepacia media from MAST Diagnostics (Bootle, United Kingdom), Lab M Ltd. (Heywood, United Kingdom), and Oxoid Ltd. (Basingstoke, United Kingdom), through the analysis of 142 clinical and environmental isolates representing all species within the B. cepacia complex (145). BCSA and MAST B. cepacia medium supported the growth of B. cepacia complex isolates most efficiently. The latter two media were also compared in a study to evaluate the sensitivities and specificities for the isolation of B. cepacia complex species from sputum specimens from CF patients

43. Miscellaneous Gram-Negative Bacteria n

797

not available. An enrichment broth consisting of Ashdown medium supplemented with 50 mg of colistin is superior to standard enrichment broth such as tryptic soy broth and increases recovery of B. pseudomallei from clinical specimens taken from colonized sites compared with plating on Ashdown medium alone (127). Selective broth cultures should be subcultured to Ashdown medium after 48 h of incubation in air at 37°C, and all inoculated plates should be incubated at 37°C in air and examined daily for 4 days before being discarded, since some colonies become apparent to the naked eye only after extended incubation. The use of selective media increases the isolation rates of S. maltophilia from clinical and environmental samples (150). Denton et al. (150) studied the sensitivity of a selective medium incorporating vancomycin, imipenem, and amphotericin B as selective agents (VIA medium) for isolating S. maltophilia from sputum samples collected from children with CF. This study compared the use of VIA medium to an existing in-house method that utilized an imipenem disk placed upon bacitracin-chocolate agar (BC medium) and reported an improved detection using VIA as a selective medium.

IDENTIFICATION B. cepacia Complex and B. gladioli

FIGURE 2 (a) B. pseudomallei colonies on MacConkey agar; (b) B. pseudomallei colonies on blood agar; (c) B. pseudomallei colonies on Ashdown medium agar. doi:10.1128/9781555817381.ch43.f2

(146). BCSA was reported as being as sensitive as MAST agar but more selective. Ashdown medium is effective for the isolation of B. pseudomallei (Fig. 2); crystal violet and gentamicin act as selective agents. It has been shown to be superior to MacConkey agar or MacConkey agar supplemented with colistin for the recovery of B. pseudomallei from clinical specimens containing mixed bacterial microbiota, such as throat, rectal, and sputum specimens (147). A more recently described selective agar, BPSA (for B. pseudomallei selective agar), was reported to improve the recovery of B. pseudomallei over that with other media (148); however, this medium is not commercially available. BPSA was more inhibitory to P. aeruginosa and B. cepacia complex species and made recognition of Burkholderia species easier due to their distinctive colony morphology. A clinical comparison of BPSA, Ashdown medium, and B. cepacia medium demonstrated equivalent sensitivity for all three media, but the selectivity of BPSA was lower than that for the other two media (149). B. cepacia medium is widely used and represents a good alternative when Ashdown medium is

Accurate identification of B. cepacia complex species can present a challenge (151). Historically, many commercial bacterial identification systems have not been able to reliably distinguish among the 18 species of the B. cepacia complex and often failed to differentiate these species from other closely related species such as B. gladioli and Ralstonia, Cupriavidus, and Pandoraea spp. (82, 112, 152). This has presented a significant problem for CF patients and their caregivers, as detailed in “Clinical Significance” above. The identification of B. cepacia complex species from CF sputum culture has a dramatic impact on patient management and is a cause of considerable anxiety for patients with CF (71, 153). Consequently, when Burkholderia, Ralstonia, Cupriavidus, or Pandoraea species are tentatively identified in a patient with CF by using a commercial system, the identity of the isolate should be confirmed by molecular techniques. To aid clinical microbiologists in the United States, the CF Foundation has established a B. cepacia reference laboratory, which uses a combination of phenotypic and genotypic methods (described below) to confirm the identity of suspected B. cepacia complex isolates (154). Further information concerning this and other reference laboratories with special expertise in Burkholderia can be found on the CF Foundation website (http://www.cff.org). B. cepacia complex species may require 3 days of incubation before colonies are seen on selective media. On MacConkey or Mueller-Hinton agar, these colonies may be punctate and tenacious, and on blood agar or selective medium such as BCSA, PC agar, or OFPBL agar, the colonies are smooth and slightly raised; occasional isolates are mucoid. On MacConkey agar, colonies of the B. cepacia complex frequently become dark pink to red due to oxidation of lactose after extended incubation (4 to 7 days). Most clinical isolates are nonpigmented, but on iron-containing media such as a triple sugar iron slant, many strains produce a bright yellow pigment. B. cepacia complex species have a characteristic dirt-like odor. The species of the B. cepacia complex are phenotypically very similar, making their differentiation, even with an extended panel of biochemical tests, rather difficult (Table 1)

798 n

BACTERIOLOGY

(112). Further, isolates within these species show considerable phenotypic variability, which is likely due to their unusually large genomes rich in insertion sequences and mobile elements such as plasmids, transposons, and bacteriophages (155). These features can contribute to genetic plasticity and diversity, which, when differentially expressed in isolates, result in variable biochemical phenotypic profiles. Most strains are weakly oxidase positive, although some strains of B. contaminans, B. lata, and B. pyrrocinia are oxidase negative. B. multivorans, B. pseudomultivorans, B. stabilis, and B. dolosa rarely oxidize sucrose. B. stabilis is ornithine decarboxylase positive, as are most B. cenocepacia strains, but is distinctive in that more than two-thirds of strains are o-nitrophenyl-βD-galactopyranoside (ONPG) negative. B. stabilis, B. lata, and most B. ambifaria strains show poor growth at 42°C. B. dolosa is usually lysine decarboxylase negative, whereas only approximately half of B. multivorans strains are negative. Other B. cepacia complex species are usually lysine decarboxylase positive. B. vietnamiensis and most B. anthina strains do not oxidize adonitol. B. anthina strains show a distinctive creamy morphology on BCSA, which also turns pink (i.e., alkaline) despite the ability of this species to utilize sucrose (156).

TABLE 1

Phenotypic differentiation of B. cepacia complex species from B. gladioli and Pandoraea spp. is also difficult (Table 1). Cellular fatty acid analysis is unable to differentiate B. cepacia complex species from B. gladioli (157). However, in contrast to B. cepacia complex species, most B. gladioli strains are oxidase negative, and whereas most B. cepacia complex strains oxidize maltose and lactose, B. gladioli typically oxidizes neither. Pandoraea spp. do not oxidize maltose, lactose, xylose, sucrose, or adonitol, and most are ONPG negative. B. cepacia complex species also may be difficult to differentiate from Ralstonia and Cupriavidus species. However, several of the latter species show a fast and strong oxidase reaction, whereas B. cepacia complex species produce a slow, weak-positive oxidase test. Further, in contrast to most B. cepacia complex species, Ralstonia and Cupriavidus are lysine decarboxylase negative and most often ONPG negative. The challenge in differentiating B. cepacia complex species prompted the development of molecular genetic diagnostic tests capable of identifying these species individually and distinguishing them (as a group) from biochemically similar species. DNA sequence differences in 16S and 23S rRNA genes have been used to develop species-specific PCR

Characteristics of the B. cepacia complex, B. gladioli, and Pandoraea spp.a

B. gladioli

Pandoraea spp.

B. vietnamiensis

+

+

+

+

V

V

+ + V − − −

+ + V V − Vβ

V + V − V −

V + V V − −

+ + V V − V

+ + V − − −

+ + + − − −

+ + − V − −

+ + + − − −

+ + + V − −

+ + + − − −

+ + + − − −

+ + V V − Vβ

+ + + V − −

+ + − − − −

+ + V − − −

V + + − − Vβ

+ V − V V

+ V V − −

+ + + V V V V − − − V

+ V + V + V V + − + +

V + + + V V + V V V +

V + + V V − + V V V +

V + + + + V + − V + +

+ + + V V + + − − V +

+ + + − + + − − − − +

+ + + V V V + V V V V

+ + + + + − + − − V +

+ + + V + − + − + + +

+ + + − + + V − − − +

+ + + − V V + − − − +

+ + + V + V + V − V +

+ + + V + − V V V + +

+ V V − + − + + − + V

+ + + + − V − − − + −

+ + V + − V + − − − +

− − + − + V − − V V +

− − − − − V − − − − V

B. arboris

B. ubonensis

V

B. stabilis

+

B. seminalis

+

B. pyrrocinia

+

B. anthina

B. ambifaria

B. pseudomultivorans

+

B. multivorans

V

B. metallica

+

B. latens

+

B. lata

V

B. dolosa

+

B. diffusa

+

B. cepacia

+

B. cenocepacia

+

Test

Oxidaseb + Growth: MacConkeyc + BCSAb + 42°Cc V Yellow pigmentc V Brown pigmentc − Hemolysisd Vβ Acid frome: Maltosec + Lactoseb + D-Xylosec + Sucroseb + Adonitolc + Nitrate reductionc,g V Lysine decarboxylaseb + Ornithine decarboxylasec − Esculin hydrolasec,g V Gelatinasec,g + PNPG or ONPGb,f +

B. contaminans

B. cepacia complex

a

Abbreviations and symbols: +, >90% of isolates are positive; V, 10 to 90% are positive; −, 2

0 0 0 0 0 0 0 100 >2

85 35 60 63 100 0 95 100 >2

100 43 100 100 100 0 0 4 0 2 21 23 6 0 22 38 19 29 68 48 3 5 99 96 39 0 0 0 0 0 Brown-tan, 52% yellow- 26% fluorescent, 27% yellow- Brown-tan, soluble orange, 44% yellowbrown, soluble soluble insolublee tan, soluble 0 0 0 0 0 0 0 0 0 93 0 0 0 0 0 0 0 0 0 0 0 0 0 0 1

a

Values are the percentages of strains positive. Data from reference 198. 793 isolates tested by using BactiDrop Oxidase (Remel, Dartford, United Kingdom) (LiPuma, unpublished). Oxidation-fermentation basal medium with 1% carbohydrate. d Lead acetate paper. e Pigment observed on Thayer-Martin agar. b c

that some strains were incorrectly identified as Pseudomonas species by MALDI-TOF MS. As is the case with several nonfermenting opportunistic species, an incomplete understanding of Stenotrophomonas taxonomy challenges accurate identification of clinical and environmental isolates.

Acidovorax, Brevundimonas, Delftia, and Comamonas spp. Characteristics of Acidovorax, Brevundimonas, Delftia, and Comamonas are given in Table 4. Acidovorax species, rarely encountered in clinical and environmental samples, are straight to slightly curved, Gram-negative bacilli that occur either singly or in short chains. They are oxidase positive and nonpigmented and have a single polar flagellum. Urease activity varies among strains (198). B. diminuta and B. vesicularis, infrequently encountered in clinical and environmental samples, have growth requirements for specific vitamins, including pantothenate, biotin, and cyanocobalamin. An additional growth requirement for B. diminuta is cysteine. Most strains of B. diminuta grow on MacConkey agar, while only ∼25% of B. vesicularis strains do so. On primary isolation media, B. diminuta colonies

are chalk white whereas many strains of B. vesicularis are characterized by an orange intracellular pigment. These organisms are oxidase positive, have a single polar flagellum, and weakly oxidize glucose (B. vesicularis more so than B. diminuta), and the vast majority fail to reduce nitrate to nitrite. The most reliable method for differentiating these two species is the test for esculin hydrolysis. Almost all strains of B. vesicularis (88%) are reported to hydrolyze this substrate, while B. diminuta strains rarely do (5%) (Table 4) (198). Comamonas spp. are straight to slightly curved, Gramnegative bacilli that occur singly or in pairs. The organisms are catalase and oxidase positive and have a single tuft of polar flagella. All human clinical Comamonas species reduce nitrate to nitrite. Phenotypic differentiation of C. terrigena from C. testosteroni is difficult, and as a result, isolates are typically reported as Comamonas spp. (Table 4). D. acidovorans is phenotypically similar to Comamonas. Key characteristics of the species include abilities to oxidize fructose and mannitol. One-quarter of the strains produce a fluorescent pigment, while approximately one-half of the strains may produce a soluble yellow to tan one (26, 198).

43. Miscellaneous Gram-Negative Bacteria n

TYPING SYSTEMS Several molecular genetic methods are available to assess the relatedness of isolates of these genera during nosocomial or community outbreak investigations. These methods are preferred over phenotypically based systems, which are less discriminatory and reproducible. Analysis of whole-genome macrorestriction profiles with pulsed-field gel electrophoresis (PFGE) has gained acceptance as a preferred genotyping method and has proved useful in numerous studies of Burkholderia, Ralstonia, and S. maltophilia (111, 203). The endonucleases XbaI and SpeI are most frequently used and typically yield a dozen or more DNA fragments for analysis. Care must be taken in interpreting PFGE profiles of Burkholderia species, however. These species have unusually large and dynamic multichromosome genomes that are prone to largescale alterations in content and arrangement (204). Consequently, epidemiologically irrelevant genomic polymorphisms may arise in the short term and confound outbreak investigations (203). Ribotyping, which relies on polymorphisms in and around rRNA operons, has been used to investigate the epidemiology of B. cepacia complex and B. pseudomallei (205–207). Both PFGE and ribotyping are relatively time-consuming and expensive to perform and are therefore not particularly well suited for routine analysis by clinical microbiology laboratories. A variety of PCRbased methods, including randomly amplified polymorphic DNA typing and repetitive-sequence PCR typing, offer attractive alternatives for genotyping S. maltophilia and Burkholderia, Ralstonia, and Pandoraea spp. (50, 122, 196, 208, 209). These methods are inexpensive and can provide rapid, reliable results. MLST, which assesses DNA sequence variation at several chromosomal loci, has been developed for numerous species, including the B. cepacia complex, B. pseudomallei, and B. mallei (167, 210, 211). A modification of the scheme developed for the B. cepacia complex enables MLST analysis of all species within the genus (168). This genotyping strategy provides robust, reproducible, and portable results and is quickly becoming the preferred method for investigating bacterial epidemiology, evolution, and population structure. Both repetitive-sequence PCR using a BOX A1R primer and multilocus variable-number tandem repeat analysis have been developed for B. pseudomallei to exclude a clonal outbreak (212, 213). Typing methods have not been reported for Brevundimonas, Delftia, Comamonas, or Acidovorax spp.

SEROLOGIC TESTS Of the organisms discussed in this chapter, B. pseudomallei is the only one for which serologic tests have been used clinically to diagnose the infection. The indirect hemagglutination assay, although not available commercially, is the most widely used test. It is performed by using a prepared antigen from strains of B. pseudomallei sensitized to sheep cells and includes unsensitized cells as a control. This assay can be adapted to a microtiter plate test system. The serologic tests currently in use have limited value for the diagnosis of melioidosis in persons who have lived in regions where melioidosis is endemic because the healthy indigenous population is often seropositive (56, 92, 214). Serologic testing is potentially useful in persons who do not normally reside in regions endemic for melioidosis, including returning travelers and laboratory workers following accidental laboratory exposure to B. pseudomallei (215). The interpretation of the indirect hemagglutination assay or other serologic assays is complicated by the fact that there are no validated guide-

803

lines, and different cutoff points have been used to define seroconversion following exposure and acute infection. Testing should be performed whenever possible on paired samples. Seroconversion with the development of detectable antibodies to B. pseudomallei in the second sample is supportive of exposure. A 4-fold rise in titer is commonly used to diagnose a range of infectious diseases, but this has not been validated for melioidosis and any reproducible rise between two samples should be viewed as possible evidence of exposure. A single high titer in persons from a nonendemic region with a relevant travel history who presents late after a putative exposure event and for whom paired sera may be less relevant is also suggestive of exposure. Some individuals with culture-proven melioidosis do not have detectable antibodies (216), and so a negative serologic test does not rule out exposure or melioidosis. Given the complexity of this situation, experts in the field should be consulted when serology is used to diagnose melioidosis. Several evaluations of a commercial rapid immunochromatographic test kit (Pan-Bio, Windsor, Queensland, Australia) for the detection of IgG and IgM antibodies to B. pseudomallei have been performed (217–219), but this test is not currently available.

ANTIMICROBIAL SUSCEPTIBILITIES Specific susceptibility testing interpretative criteria are not available for all of the species discussed in this chapter. For some species, such as the B. cepacia complex and S. maltophilia, interpretive criteria for disk diffusion testing are available for only a limited number of antibiotics. In general, MIC microbroth microdilution tests or Etests are preferred for this group of organisms. B. cepacia complex species are among the most antimicrobial-resistant bacteria encountered in the clinical laboratory. These species are intrinsically resistant to aminoglycoside and polymyxin antibiotics and are often resistant to β-lactam antibiotics due to inducible chromosomal βlactamases and altered penicillin-binding proteins (220). Antibiotic efflux pumps may mediate resistance to chloramphenicol, fluoroquinolones, and trimethoprim (221). Clinical strains may be susceptible to only a handful of agents, including trimethoprim-sulfamethoxazole (TMP-SMX), ceftazidime, chloramphenicol, minocycline, imipenem, meropenem, and some fluoroquinolones (222, 223). The glycylcycline antibiotic tigecycline shows highly variable activity in vitro (224). The relatively high MIC observed for some strains and the potential for discoloration of permanent teeth in children younger than 7 years of age limit the use of tigecycline in CF patients. Clinical and Laboratory Standards Institute (CLSI) interpretative criteria for disk diffusion susceptibility testing are available for ceftazidime, meropenem, minocycline, and TMP-SMX (225). Because isolates that are initially susceptible may become resistant during the course of therapy, susceptibility testing of repeat isolates may be warranted. Furthermore, strains recovered from patients with CF who have received repeated courses of antibiotic therapy are frequently resistant to all currently available antimicrobial agents (222). Combinations of antimicrobial agents may provide synergistic activity against resistant strains; however, antagonism with combinations is also observed in vitro (222). B. pseudomallei is intrinsically resistant to penicillins, aminoglycosides, and macrolides. Susceptibility testing should be performed to the antimicrobial agents commonly used to treat melioidosis, which are ceftazidime, imipenem or meropenem, amoxicillin-clavulanate, doxycycline, and

804 n BACTERIOLOGY

TMP-SMX (226). B. pseudomallei is usually susceptible to all of these agents with the exception of TMP-SMX, reported rates of resistance for which are in the order of 2% in Australia (227, 228) and 13 to 16% in northeast Thailand (229, 230). Disk diffusion testing of TMP-SMX overestimates resistance and is unreliable (228–230); acceptable alternatives include Etest, broth microdilution, and agar dilution (226). Fluoroquinolones are associated with a high rate of therapeutic failure (231) and should not be included in the test panel. Recent studies indicate that tigecycline has good activity against B. pseudomallei in vitro and is effective when combined with other agents in an animal model of B. pseudomallei infection (232, 233), but there are no clinical trial data to support its use. Current trends in the management of melioidosis involve an initial 10- to 14-day intensive therapy phase with ceftazidime or meropenem, followed by eradication therapy with TMP-SMX for at least 3 months (83, 84, 234, 235). A recent trial from Thailand supported dropping the traditional practice of adding doxycycline to TMP-SMX for the eradication phase of therapy (236). In Australia, TMP-SMX is added to ceftazidime or meropenem during the intensive phase for neurological, prostatic, cutaneous, and bone and joint melioidosis. Amoxicillin-clavulanate is recommended for eradication therapy in pregnancy and is an alternative to TMP-SMX in children (237). In critically ill patients requiring intensive care, meropenem or imipenem may be superior to ceftazidime, and granulocyte colony-stimulating factor is being used in some centers, although a study from Thailand showed no benefit (227, 238–240). From a molecular genotyping study of cases of recurrent melioidosis, relapse following antimicrobial therapy occurred in 9.7% of patients and a new infection occurred in 3.4% (95). Because of the potential role of B. mallei as a bioterrorism agent, studies have been done recently to determine the activities of a variety of agents against this species. B. mallei has a susceptibility profile similar to that of B. pseudomallei, except that B. mallei is susceptible to aminoglycosides and newer macrolides such as clarithromycin and azithromycin, whereas B. pseudomallei is resistant (241). Current recommended treatment and duration of therapy for glanders are the same as those for melioidosis. Guidelines on the management of accidental laboratory exposure to B. pseudomallei and B. mallei have been published (215, 242). S. maltophilia is intrinsically resistant to many classes of antibiotics. Resistance can also develop rapidly during infection (243). Resistance to β-lactam agents is mediated by at least two β-lactamases, one of which is zinc dependent and resistant to β-lactamase inhibitors and confers resistance to imipenem. Aminoglycoside and quinolone resistance results from mutations in outer membrane proteins. In a study of isolates recovered from patients with CF, doxycycline was the most active agent in vitro (244). TMP-SMX is usually active and is often used in combination with ticarcillinclavulanate, minocycline, or piperacillin-tazobactam (244). Other combinations that may be effective include ciprofloxacin paired with ticarcillin-clavulanate, ciprofloxacin and piperacillin-tazobactam, or doxycycline and ticarcillin-clavulanate. Tigecycline is reported to have good activity in vitro (224). CLSI interpretive criteria for disk diffusion susceptibility testing are available for minocycline, levofloxacin, and TMP-SMX (225). Many U.S. laboratories comment only on the activity of TMP-SMX but will test additional antibiotics such as minocycline, ceftazidime, ticarcillin-clavulanate, and ciprofloxacin or levofloxacin upon request.

In general, C. testosteroni is susceptible to extended- and broad-spectrum cephalosporins, carbapenems, quinolones, and TMP-SMX (245). D. acidovorans is frequently resistant to the aminoglycosides.

EVALUATION, INTERPRETATION, AND REPORTING OF RESULTS The species discussed in this chapter are found in the natural environment and may occasionally contaminate clinical specimens. Nevertheless, they are increasingly recognized as nosocomial and opportunistic pathogens, especially in certain patient populations, such as persons with CF. They are also frequently misidentified by commercial microbial identification systems. Therefore, their recovery in the clinical laboratory must be given careful consideration. In particular, species of the B. cepacia complex are not reliably differentiated by phenotypic analyses, and their recovery from persons with CF has serious consequences with respect to patient management and psychosocial well-being (69). Identification of these species should be confirmed by genotypic analyses at a reference laboratory and should promptly be reported to the CF care team. Recovery of B. pseudomallei and B. mallei in any context should always be considered to reflect clinical disease. Identification of these species should be confirmed by a reference laboratory with experience with these species. Care must be given to ensure that culture handling and shipping comply with current biosafety regulations (see chapter 18). Identification of these species must be reported to public health officials due to the potential of these species as agents of bioterrorism (see chapter 14). The relevance of the recovery of the other genera described in this chapter, outside the context of CF, is less clear and should be interpreted with caution. Interpretive criteria for disk diffusion antimicrobial susceptibility testing of most of these species are lacking; MIC broth microdilution and the Etest are therefore the preferred methodologies for susceptibility testing. For multiresistant strains, consideration could be given to testing in reference laboratories for synergy with double or triple combinations of antimicrobial agents (222). It is important to note, however, that neither checkerboard MIC broth microdilution testing nor multiple combination bactericidal antibiotic testing is standardized at present.

REFERENCES 1. Yabuuchi E, Kosako Y, Oyaizu H, Yano I, Hotta H, Hashimoto Y, Ezaki T, Arakawa M. 1992. Proposal of Burkholderia gen. nov. and transfer of seven species of the genus Pseudomonas homology group II to the new genus, with the type species Burkholderia cepacia (Palleroni and Holmes 1981) comb. nov. Microbiol Immunol 36:1251– 1275. 2. Yabuuchi E, Kosako Y, Yano I, Hotta H, Nishiuchi Y. 1995. Transfer of two Burkholderia and an Alcaligenes species to Ralstonia gen. nov.: proposal of Ralstonia pickettii (Ralston, Palleroni and Doudoroff 1973) comb. nov., Ralstonia solanacearum (Smith 1896) comb. nov. and Ralstonia eutropha (Davis 1969) comb. nov. Microbiol Immunol 39:897– 904. 3. Coenye T, Falsen E, Hoste B, Ohlén M, Goris J, Govan JR, Gillis M, Vandamme P. 2000. Description of Pandoraea gen. nov. with Pandoraea apista sp. nov., Pandoraea pulmonicola sp. nov., Pandoraea pnomenusa sp. nov., Pandoraea sputorum sp. nov. and Pandoraea norimbergensis comb. nov. Int J Syst Evol Microbiol 50:887–899. 4. Vaneechoutte M, Kampfer P, De Baere T, Falsen E, Verschraegen G. 2004. Wautersia gen. nov., a novel genus

43. Miscellaneous Gram-Negative Bacteria n

5.

6.

7.

8.

9.

10.

11.

12.

13.

14.

15.

16.

17.

18.

19.

accommodating the phylogenetic lineage including Ralstonia eutropha and related species, and proposal of Ralstonia [Pseudomonas] syzygii (Roberts et al. 1990) comb. nov. Int J Syst Evol Microbiol 54:317–327. Vandamme P, Coenye T. 2004. Taxonomy of the genus Cupriavidus: a tale of lost and found. Int J Syst Evol Microbiol 54:2285–2289. Makkar NS, Casida LE. 1987. Cupriavidus necator gen. nov., sp. nov.: a nonobligate bacterial predator of bacteria in soil. Int J Syst Bacteriol 37:323–326. Sneath PHA. 1992. International Code of Nomenclature of Bacteria. 1990 revision. American Society for Microbiology, Washington, DC. Coenye T, Holmes B, Kersters K, Govan JR, Vandamme P. 1999. Burkholderia cocovenenans (van Damme et al. 1960) Gillis et al. 1995 and Burkholderia vandii Urakami et al. 1994 are junior synonyms of Burkholderia gladioli (Severini 1913) Yabuuchi et al. 1993 and Burkholderia plantarii (Azegami et al. 1987) Urakami et al. 1994, respectively. Int J Syst Bacteriol 49:37–42. Vandamme P, Dawyndt P. 2011. Classification and identification of the Burkholderia cepacia complex: past, present and future. Syst Appl Microbiol 34:87–95. Peeters C, Zlosnik JE, Spilker T, Hird TJ, LiPuma JJ, Vandamme P. 2013. Burkholderia pseudomultivorans sp. nov., a novel Burkholderia cepacia complex species from human respiratory samples and the rhizosphere. Syst Appl Microbiol 36:483–489. Reik R, Spilker T, LiPuma JJ. 2005. Distribution of Burkholderia cepacia complex species among isolates recovered from persons with or without cystic fibrosis. J Clin Microbiol 43:2926–2928. Coenye T, Laevens S, Willems A, Ohlén M, Hannant W, Govan JR, Gillis M, Falsen E, Vandamme P. 2001. Burkholderia fungorum sp. nov. and Burkholderia caledonica sp. nov., two new species isolated from the environment, animals and human clinical samples. Int J Syst Evol Microbiol 51:1099–1107. Weinberg JB, Alexander BD, Majure JM, Williams LW, Kim JY, Vandamme P, LiPuma JJ. 2007. Burkholderia glumae infection in an infant with chronic granulomatous disease. J Clin Microbiol 45:662–665. Deris ZZ, Van Rostenberghe H, Habsah H, Noraida R, Tan GC, Chan YY, Rosliza AR, Ravichandran M. 2010. First isolation of Burkholderia tropica from a neonatal patient successfully treated with imipenem. Int J Infect Dis 14:e73– e74. Gee JE, Glass MB, Lackner G, Helsel LO, Daneshvar M, Hollis DG, Jordan J, Morey R, Steigerwalt A, Hertweck C. 2011. Characterization of Burkholderia rhizoxinica and B. endofungorum isolated from clinical specimens. PLoS One 6:e15731. doi:10.1371/journal.pone.0015731. Gerrits GP, Klaassen C, Coenye T, Vandamme P, Meis JF. 2005. Burkholderia fungorum septicemia. Emerg Infect Dis 11:1115–1117. De Baere T, Steyaert S, Wauters G, Des Vos P, Goris J, Coenye T, Suyama T, Verschraegen G, Vaneechoutte MS. 2001. Classification of Ralstonia pickettii biovar 3/‘thomasii’ strains (Pickett 1994) and of new isolates related to nosocomial recurrent meningitis as Ralstonia mannitolytica sp. nov. Int J Syst Evol Microbiol 51:547–558. Coenye T, Goris J, De Vos P, Vandamme P, LiPuma JJ. 2003. Classification of Ralstonia pickettii-like isolates from the environment and clinical samples as Ralstonia insidiosa sp. nov. Int J Syst Evol Microbiol 53:1075–1080. Vandamme P, Goris J, Coenye T, Hoste B, Janssens D, Kersters K, De Vos P, Falsen E. 1999. Assignment of Centers for Disease Control group IVc-2 to the genus Ralstonia as Ralstonia paucula sp. nov. Int J Syst Bacteriol 49:663–669.

805

20. Coenye T, Falsen E, Vancanneyt M, Hoste B, Govan JR, Kersters K, Vandamme P. 1999. Classification of Alcaligenes faecalis-like isolates from the environment and human clinical samples as Ralstonia gilardii sp. nov. Int J Syst Bacteriol 49:405–413. 21. Coenye T, Vandamme P, LiPuma JJ. 2003. Ralstonia respiraculi sp. nov., isolated from the respiratory tract of cystic fibrosis patients. Int J Syst Evol Microbiol 53:1339–1342. 22. Chen WM, Laevens S, Lee TM, Coenye T, De Vos P, Mergeay M, Vandamme P. 2001. Ralstonia taiwanensis sp. nov., isolated from root nodules of Mimosa species and sputum of a cystic fibrosis patient. Int J Syst Evol Microbiol 51:1729–1735. 23. Sahin N, Tani A, Kotan R, Sedlacek I, Kimbara K, Tamer AU. 2011. Pandoraea oxalativorans sp. nov., Pandoraea faecigallinarum sp. nov. and Pandoraea vervacti sp. nov., isolated from oxalate-enriched culture. Int J Syst Evol Microbiol 61:2247–2253. 24. Anandham R, Indiragandhi P, Kwon SW, Sa TM, Jeon CO, Kim YK, Jee HJ. 2010. Pandoraea thiooxydans sp. nov., a facultatively chemolithotrophic, thiosulfate-oxidizing bacterium isolated from rhizosphere soils of sesame (Sesamum indicum L.). Int J Syst Evol Microbiol 60:21–26. 25. Henry D, Campbell M, McGimpsey C, Clarke A, Louden L, Burns JL, Roe MH, Vandamme P, Speert D. 1999. Comparison of isolation media for recovery of Burkholderia cepacia complex from respiratory secretions of patients with cystic fibrosis. J Clin Microbiol 37:1004–1007. 26. Willems A, De Ley J, Gillis M, Kersters K. 1991. Comamonadaceae, a new family encompassing the acidovorans rRNA complex, including Variovorax paradoxus gen. nov., comb nov., for Alcaligenes paradoxus (Davis 1969). Int J Syst Evol Microbiol 41:445–450. 27. Wen A, Fegan M, Hayward C, Chakraborty S, Sly LI. 1999. Phylogenetic relationships among members of the Comamonadaceae, and description of Delftia acidovorans (den Dooren de Jong 1926 and Tamaoka et al. 1987) gen. nov., comb. nov. Int J Syst Bacteriol 49:567–576. 28. Willems A, Pot B, Falsen E, Vandamme P, Gillis M, Kersters K, De Ley J. 1991. Polyphasic taxonomic study of the emended genus Comamonas: relationship to Aquaspirillum aquaticum, E. Falsen group 10, and other clinical isolates. Int J Syst Evol Microbiol 41:427–444. 29. Wauters G, De Baere T, Willems A, Falsen E, Vaneechoutte M. 2003. Description of Comamonas aquatica comb. nov. and Comamonas kerstersii sp. nov. for two subgroups of Comamonas terrigena and emended description of Comamonas terrigena. Int J Syst Evol Microbiol 53:859– 862. 30. Chen WM, Lin YS, Sheu DS, Sheu SY. 2012. Delftia litopenaei sp. nov., a poly-β-hydroxybutyrate-accumulating bacterium isolated from a freshwater shrimp culture pond. Int J Syst Evol Microbiol 62:2315–2321. 31. Gardan L, Stead DE, Dauga C, Gillis M. 2003. Acidovorax valerianellae sp. nov., a novel pathogen of lamb’s lettuce [Valerianella locusta (L.) Laterr]. Int J Syst Evol Microbiol 53:795–800. 32. Vaneechoutte M, Janssens M, Avesani V, Delmee M, Deschaght P. 2013. Description of Acidovorax wautersii sp. nov. to accommodate clinical isolates and an environmental isolate, most closely related to Acidovorax avenae. Int J Syst Evol Microbiol 63:2203–2206. 33. Segers P, Vancanneyt M, Pot B, Torck U, Hoste B, Dewettinck D, Falsen E, Kersters K, De Vos P. 1994. Classification of Pseudomonas diminuta Leifson and Hugh 1954 and Pseudomonas vesicularis Busing, Doll, and Freytag 1953 in Brevundimonas gen. nov. as Brevundimonas diminuta comb. nov. and Brevundimonas vesicularis comb. nov., respectively. Int J Syst Bacteriol 44:499–510.

806 n

BACTERIOLOGY

34. Estrela AB, Abraham WR. 2010. Brevundimonas vancanneytii sp. nov., isolated from blood of a patient with endocarditis. Int J Syst Evol Microbiol 60:2129–2134. 35. Palleroni NJ, Bradbury JF. 1993. Stenotrophomonas, a new bacterial genus for Xanthomonas maltophilia (Hugh 1980) Swings et al. 1983. Int J Syst Bacteriol 43:606–609. 36. Swings J, De Vos P, Van den Mooter M, De Ley J. 1983. Transfer of Pseudomonas maltophilia Hugh 1981 to the genus Xanthomonas as Xanthomonas maltophilia (Hugh 1981) comb. nov. Int J Syst Bacteriol 33:409–413. 37. Brooke JS. 2012. Stenotrophomonas maltophilia: an emerging global opportunistic pathogen. Clin Microbiol Rev 25:2–41. 38. Vasileuskaya-Schulz Z, Kaiser S, Maier T, Kostrzewa M, Jonas D. 2011. Delineation of Stenotrophomonas spp. by multi-locus sequence analysis and MALDI-TOF mass spectrometry. Syst Appl Microbiol 34:35–39. 39. Holt J, Krieg NR, Sneath PHA, Staley JT, Williams ST. 1994. Bergey’s Manual of Determinative Bacteriology, 9th ed, vol 1. Williams and Wilkins, Philadelphia, PA. 40. Palleroni NJ. 1984. Genus I. Pseudomonas Migula 1984, 237AL, p 141–199. In Holt J, Krieg NR (ed), Bergey’s Manual of Systematic Bacteriology, vol 1. Williams and Wilkins, Baltimore, MD. 41. LiPuma JJ. 2010. The changing microbial epidemiology in cystic fibrosis. Clin Microbiol Rev 23:299–323. 42. Parke JL, Gurian-Sherman D. 2001. Diversity of the Burkholderia cepacia complex and implications for risk assessment of biological control strains. Annu Rev Phytopathol 39:225–258. 43. Moore JE, Xu J, Millar BC, Crowe M, Elborn JS. 2002. Improved molecular detection of Burkholderia cepacia genomovar III and Burkholderia multivorans directly from sputum of patients with cystic fibrosis. J Microbiol Methods 49:183–191. 44. Ramette A, LiPuma JJ, Tiedje JM. 2005. Species abundance and diversity of Burkholderia cepacia complex in the environment. Appl Environ Microbiol 71:1193–1201. 45. Moore JE, McIlhatton B, Shaw A, Murphy PG, Elborn JS. 2001. Occurrence of Burkholderia cepacia in foods and waters: clinical implications for patients with cystic fibrosis. J Food Prot 64:1076–1078. 46. Jimenez L. 2007. Microbial diversity in pharmaceutical product recalls and environments. PDA J Pharm Sci Technol 61:383–399. 47. Torbeck L, Raccasi D, Guilfoyle DE, Friedman RL, Hussong D. 2011. Burkholderia cepacia: this decision is overdue. PDA J Pharm Sci Technol 65:535–543. 48. Johnson WM, Tyler SD, Rozee KR. 1994. Linkage analysis of geographic and clinical clusters in Pseudomonas cepacia infections by multilocus enzyme electrophoresis and ribotyping. J Clin Microbiol 32:924–930. 49. Pitt TL, Kaufmann ME, Patel PS, Benge LC, Gaskin S, Livermore DM. 1996. Type characterisation and antibiotic susceptibility of Burkholderia (Pseudomonas) cepacia isolates from patients with cystic fibrosis in the United Kingdom and the Republic of Ireland. J Med Microbiol 44:203–210. 50. Chen JS, Witzmann KA, Spilker T, Fink RJ, LiPuma JJ. 2001. Endemicity and inter-city spread of Burkholderia cepacia genomovar III in cystic fibrosis. J Pediatr 139:643– 649. 51. Coenye T, Spilker T, Van Schoor A, LiPuma JJ, Vandamme P. 2004. Recovery of Burkholderia cenocepacia strain PHDC from cystic fibrosis patients in Europe. Thorax 59:952–954. 52. LiPuma JJ, Spilker T, Coenye T, Gonzalez CF. 2002. An epidemic Burkholderia cepacia complex strain identified in soil. Lancet 359:2002–2003. 53. Coenye T, LiPuma JJ. 2003. Population structure analysis of Burkholderia cepacia genomovar III: varying degrees of genetic recombination characterize major clonal complexes. Microbiology 149:77–88.

54. Chantratita N, Wuthiekanun V, Limmathurotsakul D, Vesaratchavest M, Thanwisai A, Amornchai P, Tumapa S, Feil EJ, Day NP, Peacock SJ. 2008. Genetic diversity and microevolution of Burkholderia pseudomallei in the environment. PLoS Negl Trop Dis 2:e182. doi:10.1371/journal.pntd.0000182. 55. Parry CM, Wuthiekanun V, Hoa NT, Diep TS, Thao LT, Loc PV, Wills BA, Wain J, Hien TT, White NJ, Farrar JJ. 1999. Melioidosis in southern Vietnam: clinical surveillance and environmental sampling. Clin Infect Dis 29:1323–1326. 56. Wuthiekanun V, Pheaktra N, Putchhat H, Sin L, Sen B, Kumar V, Langla S, Peacock SJ, Day NP. 2008. Burkholderia pseudomallei antibodies in children, Cambodia. Emerg Infect Dis 14:301–303. 57. Kaestli M, Mayo M, Harrington G, Ward L, Watt F, Hill JV, Cheng AC, Currie BJ. 2009. Landscape changes influence the occurrence of the melioidosis bacterium Burkholderia pseudomallei in soil in northern Australia. PLoS Negl Trop Dis 3:e364. doi:10.1371/journal.pntd.0000364. 58. Kaestli M, Schmid M, Mayo M, Rothballer M, Harrington G, Richardson L, Hill A, Hill J, Tuanyok A, Keim P, Hartmann A, Currie BJ. 2012. Out of the ground: aerial and exotic habitats of the melioidosis bacterium Burkholderia pseudomallei in grasses in Australia. Environ Microbiol 14:2058–2070. 59. Gee JE, Glass MB, Novak RT, Gal D, Mayo MJ, Steigerwalt AG, Wilkins PP, Currie BJ. 2008. Recovery of a Burkholderia thailandensis-like isolate from an Australian water source. BMC Microbiol 8:54. doi:10.1186/1471-21808-54. 60. Currie BJ, Dance DA, Cheng AC. 2008. The global distribution of Burkholderia pseudomallei and melioidosis: an update. Trans R Soc Trop Med Hyg 102(Suppl 1):S1–S4. 61. Limmathurotsakul D, Dance DA, Wuthiekanun V, Kaestli M, Mayo M, Warner J, Wagner DM, Tuanyok A, Wertheim H, Yoke Cheng T, Mukhopadhyay C, Puthucheary S, Day NP, Steinmetz I, Currie BJ, Peacock SJ. 2013. Systematic review and consensus guidelines for environmental sampling of Burkholderia pseudomallei. PLoS Negl Trop Dis 7:e2105. doi:10.1371/journal.pntd.0002105. 62. Limmathurotsakul D, Wongratanacheewin S, Teerawattanasook N, Wongsuvan G, Chaisuksant S, Chetchotisakd P, Chaowagul W, Day NP, Peacock SJ. 2010. Increasing incidence of human melioidosis in northeast Thailand. Am J Trop Med Hyg 82:1113–1117. 63. Dance DA, Smith MD, Aucken HM, Pitt TL. 1999. Imported melioidosis in England and Wales. Lancet 353:208. 64. Klausner JD, Zukerman C, Limaye AP, Corey L. 1999. Outbreak of Stenotrophomonas maltophilia bacteremia among patients undergoing bone marrow transplantation: association with faulty replacement of handwashing soap. Infect Control Hosp Epidemiol 20:756–758. 65. Weber DJ, Rutala WA, Blanchet CN, Jordan M, Gergen MF. 1999. Faucet aerators: a source of patient colonization with Stenotrophomonas maltophilia. Am J Infect Control 27:59–63. 66. Greenberg DE, Goldberg JB, Stock F, Murray PR, Holland SM, Lipuma JJ. 2009. Recurrent Burkholderia infection in patients with chronic granulomatous disease: 11year experience at a large referral center. Clin Infect Dis 48:1577–1579. 67. Isles A, Maclusky I, Corey M, Gold R, Prober C, Fleming P, Levison H. 1984. Pseudomonas cepacia infection in cystic fibrosis: an emerging problem. J Pediatr 104:206–210. 68. Liou TG, Adler FR, Fitzsimmons SC, Cahill BC, Hibbs JR, Marshall BC. 2001. Predictive 5-year survivorship model of cystic fibrosis. Am J Epidemiol 153:345–352. 69. LiPuma JJ. 2003. Burkholderia and emerging pathogens in cystic fibrosis. Semin Respir Crit Care Med 24:681–692.

43. Miscellaneous Gram-Negative Bacteria n 70. Cystic Fibrosis Foundation. 2012. Cystic Fibrosis Foundation Patient Registry. 2011 Annual Data Report. Cystic Fibrosis Foundation, Bethesda, MD. http://www.cff.org/ UploadedFiles/research/ClinicalResearch/2011-PatientRegistry.pdf. 71. LiPuma JJ. 2001. Burkholderia cepacia complex: a contraindication to lung transplantation in cystic fibrosis? Transpl Infect Dis 3:149–160. 72. Govan JR, Brown AR, Jones AM. 2007. Evolving epidemiology of Pseudomonas aeruginosa and the Burkholderia cepacia complex in cystic fibrosis lung infection. Future Microbiol 2:153–164. 73. Alexander BD, Petzold EW, Reller LB, Palmer SM, Davis RD, Woods CW, LiPuma JJ. 2008. Survival after lung transplantation of cystic fibrosis patients infected with Burkholderia cepacia complex. Am J Transplant 8:1025–1030. 74. Aris RM, Routh JC, LiPuma JJ, Heath DG, Gilligan PH. 2001. Lung transplantation for cystic fibrosis patients with Burkholderia cepacia complex. Survival linked to genomovar type. Am J Respir Crit Care Med 164:2102–2106. 75. Murray S, Charbeneau J, Marshall BC, LiPuma JJ. 2008. Impact of Burkholderia infection on lung transplantation in cystic fibrosis. Am J Respir Crit Care Med 178:363–371. 76. Blackburn L, Brownlee K, Conway S, Denton M. 2004. ‘Cepacia syndrome’ with Burkholderia multivorans, 9 years after initial colonization. J Cyst Fibros 3:133–134. 77. Otag F, Ersoz G, Salcioglu M, Bal C, Schneider I, Bauernfeind A. 2005. Nosocomial bloodstream infections with Burkholderia stabilis. J Hosp Infect 59:46–52. 78. Ledson MJ, Gallagher MJ, Jackson M, Hart CA, Walshaw MJ. 2002. Outcome of Burkholderia cepacia colonisation in an adult cystic fibrosis centre. Thorax 57:142–145. 79. Kalish LA, Waltz DA, Dovey M, Potter-Bynoe G, McAdam AJ, Lipuma JJ, Gerard C, Goldmann D. 2006. Impact of Burkholderia dolosa on lung function and survival in cystic fibrosis. Am J Respir Crit Care Med 173:421–425. 80. Graves M, Robin T, Chipman AM, Wong J, Khashe S, Janda JM. 1997. Four additional cases of Burkholderia gladioli infection with microbiological correlates and review. Clin Infect Dis 25:838–842. 81. Jones AM, Stanbridge TN, Isalska BJ, Dodd ME, Webb AK. 2001. Burkholderia gladioli: recurrent abscesses in a patient with cystic fibrosis. J Infect 42:69–71. 82. Shelly DB, Spilker T, Gracely EJ, Coenye T, Vandamme P, LiPuma JJ. 2000. Utility of commercial systems for identification of Burkholderia cepacia complex from cystic fibrosis sputum culture. J Clin Microbiol 38:3112–3115. 83. Cheng AC, Currie BJ. 2005. Melioidosis: epidemiology, pathophysiology, and management. Clin Microbiol Rev 18:383–416. 84. Wiersinga WJ, Currie BJ, Peacock SJ. 2012. Melioidosis. N Engl J Med 367:1035–1044. 85. Parameswaran U, Baird RW, Ward LM, Currie BJ. 2012. Melioidosis at Royal Darwin Hospital in the big 2009– 2010 wet season: comparison with the preceding 20 years. Med J Aust 196:345–348. 86. Athan E, Allworth AM, Engler C, Bastian I, Cheng AC. 2005. Melioidosis in tsunami survivors. Emerg Infect Dis 11:1638–1639. 87. Chierakul W, Winothai W, Wattanawaitunechai C, Wuthiekanun V, Rugtaengan T, Rattanalertnavee J, Jitpratoom P, Chaowagul W, Singhasivanon P, White NJ, Day NP, Peacock SJ. 2005. Melioidosis in 6 tsunami survivors in southern Thailand. Clin Infect Dis 41:982–990. 88. O’Carroll MR, Kidd TJ, Coulter C, Smith HV, Rose BR, Harbour C, Bell SC. 2003. Burkholderia pseudomallei: another emerging pathogen in cystic fibrosis. Thorax 58:1087–1091. 89. Visca P, Cazzola G, Petrucca A, Braggion C. 2001. Travel-associated Burkholderia pseudomallei infection (meli-

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

100.

101.

102.

103.

104.

105.

807

oidosis) in a patient with cystic fibrosis: a case report. Clin Infect Dis 32:E15–E16. Limmathurotsakul D, Kanoksil M, Wuthiekanun V, Kitphati R, deStavola B, Day NP, Peacock SJ. 2013. Activities of daily living associated with acquisition of melioidosis in northeast Thailand: a matched case-control study. PLoS Negl Trop Dis 7:e2072. doi:10.1371/journal.pntd.0002072. Currie BJ, Jacups SP. 2003. Intensity of rainfall and severity of melioidosis, Australia. Emerg Infect Dis 9:1538– 1542. Wuthiekanun V, Chierakul W, Langa S, Chaowagul W, Panpitpat C, Saipan P, Thoujaikong T, Day NP, Peacock SJ. 2006. Development of antibodies to Burkholderia pseudomallei during childhood in melioidosis-endemic northeast Thailand. Am J Trop Med Hyg 74:1074– 1075. Ngauy V, Lemeshev Y, Sadkowski L, Crawford G. 2005. Cutaneous melioidosis in a man who was taken as a prisoner of war by the Japanese during World War II. J Clin Microbiol 43:970–972. Currie BJ, Jacups SP, Cheng AC, Fisher DA, Anstey NM, Huffam SE, Krause VL. 2004. Melioidosis epidemiology and risk factors from a prospective whole-population study in northern Australia. Trop Med Int Health 9:1167–1174. Limmathurotsakul D, Chaowagul W, Chierakul W, Stepniewska K, Maharjan B, Wuthiekanun V, White NJ, Day NP, Peacock SJ. 2006. Risk factors for recurrent melioidosis in northeast Thailand. Clin Infect Dis 43:979– 986. Currie BJ, Ward L, Cheng AC. 2010. The epidemiology and clinical spectrum of melioidosis: 540 cases from the 20 year Darwin prospective study. PLoS Negl Trop Dis 4:e900. doi:10.1371/journal.pntd.0000900. Stoesser N, Pocock J, Moore CE, Soeng S, Chhat HP, Sar P, Limmathurotsakul D, Day N, Thy V, Sar V, Parry CM. 2012. Pediatric suppurative parotitis in Cambodia between 2007 and 2011. Pediatr Infect Dis J 31:865– 868. Meumann EM, Cheng AC, Ward L, Currie BJ. 2012. Clinical features and epidemiology of melioidosis pneumonia: results from a 21-year study and review of the literature. Clin Infect Dis 54:362–369. Gibney KB, Cheng AC, Currie BJ. 2008. Cutaneous melioidosis in the tropical top end of Australia: a prospective study and review of the literature. Clin Infect Dis 47:603–609. Morse LP, Moller CC, Harvey E, Ward L, Cheng AC, Carson PJ, Currie BJ. 2009. Prostatic abscess due to Burkholderia pseudomallei: 81 cases from a 19-year prospective melioidosis study. J Urol 182:542–547; discussion 547. Centers for Disease Control and Prevention (CDC). 2000. Laboratory-acquired human glanders—Maryland, May 2000. MMWR Morb Mortal Wkly Rep 49:532–535. Agger WA, Cogbill TH, Busch H Jr, Landercasper J, Callister SM. 1986. Wounds caused by corn-harvesting machines: an unusual source of infection due to gramnegative bacilli. Rev Infect Dis 8:927–931. Hanes SD, Demirkan K, Tolley E, Boucher BA, Croce MA, Wood GC, Fabian TC. 2002. Risk factors for lateonset nosocomial pneumonia caused by Stenotrophomonas maltophilia in critically ill trauma patients. Clin Infect Dis 35:228–235. Denton M, Kerr KG. 1998. Microbiological and clinical aspects of infection associated with Stenotrophomonas maltophilia. Clin Microbiol Rev 11:57–80. Sattler CA, Mason EO Jr, Kaplan SL. 2000. Nonrespiratory Stenotrophomonas maltophilia infection at a children’s hospital. Clin Infect Dis 31:1321–1330.

808 n

BACTERIOLOGY

106. Razvi S, Quittell L, Sewall A, Quinton H, Marshall B, Saiman L. 2009. Respiratory microbiology of patients with cystic fibrosis in the United States, 1995 to 2005. Chest 136:1554–1560. 107. Graff GR, Burns JL. 2002. Factors affecting the incidence of Stenotrophomonas maltophilia isolation in cystic fibrosis. Chest 121:1754–1760. 108. Goss CH, Otto K, Aitken ML, Rubenfeld GD. 2002. Detecting Stenotrophomonas maltophilia does not reduce survival of patients with cystic fibrosis. Am J Respir Crit Care Med 166:356–361. 109. Wertheim WA, Markovitz DM. 1992. Osteomyelitis and intervertebral discitis caused by Pseudomonas pickettii. J Clin Microbiol 30:2506–2508. 110. Boutros N, Gonullu N, Casetta A, Guibert M, Ingrand D, Lebrun L. 2002. Ralstonia pickettii traced in blood culture bottles. J Clin Microbiol 40:2666–2667. 111. Chetoui H, Melin P, Struelens MJ, Delhalle E, Nigo MM, De Ryck R, De Mol P. 1997. Comparison of biotyping, ribotyping, and pulsed-field gel electrophoresis for investigation of a common-source outbreak of Burkholderia pickettii bacteremia. J Clin Microbiol 35:1398–1403. 112. Henry DA, Mahenthiralingam E, Vandamme P, Coenye T, Speert DP. 2001. Phenotypic methods for determining genomovar status of the Burkholderia cepacia complex. J Clin Microbiol 39:1073–1078. 113. Coenye T, Spilker T, Reik R, Vandamme P, LiPuma JJ. 2005. Use of PCR analyses to define the distribution of Ralstonia species recovered from patients with cystic fibrosis. J Clin Microbiol 43:3463–3466. 114. Chi CY, Fung CP, Wong WW, Liu CY. 2004. Brevundimonas bacteremia: two case reports and literature review. Scand J Infect Dis 36:59–61. 115. Gilad J, Borer A, Peled N, Riesenberg K, Tager S, Appelbaum A, Schlaeffer F. 2000. Hospital-acquired Brevundimonas vesicularis septicaemia following openheart surgery: case report and literature review. Scand J Infect Dis 32:90–91. 116. Han XY, Andrade RA. 2005. Brevundimonas diminuta infections and its resistance to fluoroquinolones. J Antimicrob Chemother 55:853–859. 117. Arda B, Aydemir S, Yamazhan T, Hassan A, Tünger A, Serter D. 2003. Comamonas testosteroni meningitis in a patient with recurrent cholesteatoma. APMIS 111:474– 476. 118. Cooper GR, Staples ED, Iczkowski KA, Clancy CJ. 2005. Comamonas (Pseudomonas) testosteroni endocarditis. Cardiovasc Pathol 14:145–149. 119. Le Moal G, Paccalin M, Breux JP, Roblot F, Roblot P, Becq-Giraudon B. 2001. Central venous catheter-related infection due to Comamonas testosteroni in a woman with breast cancer. Scand J Infect Dis 33:627–628. 120. Ender PT, Dooley DP, Moore RH. 1996. Vascular catheter-related Comamonas acidovorans bacteremia managed with preservation of the catheter. Pediatr Infect Dis J 15:918–920. 121. Xu J, Moore JE, Millar BC, Alexander HD, McClurg R, Morris TC, Rooney PJ. 2004. Improved laboratory diagnosis of bacterial and fungal infections in patients with hematological malignancies using PCR and ribosomal RNA sequence analysis. Leuk Lymphoma 45:1637– 1641. 122. Coenye T, Goris J, Spilker T, Vandamme P, LiPuma JJ. 2002. Characterization of unusual bacteria isolated from respiratory secretions of cystic fibrosis patients and description of Inquilinus limosus gen. nov., sp. nov. J Clin Microbiol 40:2062–2069. 123. Johnson LN, Han JY, Moskowitz SM, Burns JL, Qin X, Englund JA. 2004. Pandoraea bacteremia in a cystic fibrosis patient with associated systemic illness. Pediatr Infect Dis J 23:881–882.

124. Jørgensen IM, Johansen HK, Frederiksen B, Pressler T, Hansen A, Vandamme P, Høiby N, Koch C. 2003. Epidemic spread of Pandoraea apista, a new pathogen causing severe lung disease in cystic fibrosis patients. Pediatr Pulmonol 36:439–446. 125. Daneshvar MI, Hollis DG, Steigerwalt AG, Whitney AM, Spangler L, Douglas MP, Jordan JG, MacGregor JP, Hill BC, Tenover FC, Brenner DJ, Weyant RS. 2001. Assignment of CDC weak oxidizer group 2 (WO2) to the genus Pandoraea and characterization of three new Pandoraea genomospecies. J Clin Microbiol 39:1819– 1826. 126. Stryjewski ME, LiPuma JJ, Messier RH Jr, Reller LB, Alexander BD. 2003. Sepsis, multiple organ failure, and death due to Pandoraea pnomenusa infection after lung transplantation. J Clin Microbiol 41:2255–2257. 127. Cheng AC, Wuthiekanun V, Limmathurosakul D, Wongsuvan G, Day NP, Peacock SJ. 2006. Role of selective and nonselective media for isolation of Burkholderia pseudomallei from throat swabs of patients with melioidosis. J Clin Microbiol 44:2316. 128. McDowell A, Mahenthiralingam E, Moore JE, Dunbar KE, Webb AK, Dodd ME, Martin SL, Millar BC, Scott CJ, Crowe M, Elborn JS. 2001. PCR-based detection and identification of Burkholderia cepacia complex pathogens in sputum from cystic fibrosis patients. J Clin Microbiol 39:4247–4255. 129. Meumann EM, Novak RT, Gal D, Kaestli ME, Mayo M, Hanson JP, Spencer E, Glass MB, Gee JE, Wilkins PP, Currie BJ. 2006. Clinical evaluation of a type III secretion system real-time PCR assay for diagnosing melioidosis. J Clin Microbiol 44:3028–3030. 130. Desakorn V, Smith MD, Wuthiekanun V, Dance DA, Aucken H, Suntharasamai P, Rajchanuwong A, White NJ. 1994. Detection of Pseudomonas pseudomallei antigen in urine for the diagnosis of melioidosis. Am J Trop Med Hyg 51:627–633. 131. Smith MD, Wuthiekanun V, Walsh AL, Teerawattanasook N, Desakorn V, Suputtamongkol Y, Pitt TL, White NJ. 1995. Latex agglutination for rapid detection of Pseudomonas pseudomallei antigen in urine of patients with melioidosis. J Clin Pathol 48:174–176. 132. Wuthiekanun V, Desakorn V, Wongsuvan G, Amornchai P, Cheng AC, Maharjan B, Limmathurotsakul D, Chierakul W, White NJ, Day NP, Peacock SJ. 2005. Rapid immunofluorescence microscopy for diagnosis of melioidosis. Clin Diagn Lab Immunol 12:555–556. 133. Tandhavanant S, Wongsuvan G, Wuthiekanun V, Teerawattanasook N, Day NP, Limmathurotsakul D, Peacock SJ, Chantratita N. 2013. Monoclonal antibodybased immunofluorescence microscopy for the rapid identification of Burkholderia pseudomallei in clinical specimens. Am J Trop Med Hyg 89:165–168. 134. Haase A, Brennan M, Barrett S, Wood Y, Huffam S, O’Brien D, Currie B. 1998. Evaluation of PCR for diagnosis of melioidosis. J Clin Microbiol 36:1039–1041. 135. Novak RT, Glass MB, Gee JE, Gal D, Mayo MJ, Currie BJ, Wilkins PP. 2006. Development and evaluation of a real-time PCR assay targeting the type III secretion system of Burkholderia pseudomallei. J Clin Microbiol 44:85– 90. 136. Thibault FM, Valade E, Vidal DR. 2004. Identification and discrimination of Burkholderia pseudomallei, B. mallei, and B. thailandensis by real-time PCR targeting type III secretion system genes. J Clin Microbiol 42:5871–5874. 137. Tomaso H, Pitt TL, Landt O, Al Dahouk S, Scholz HC, Reisinger EC, Sprague LD, Rathmann I, Neubauer H. 2005. Rapid presumptive identification of Burkholderia pseudomallei with real-time PCR assays using fluorescent hybridization probes. Mol Cell Probes 19:9–20.

43. Miscellaneous Gram-Negative Bacteria n 138. Chantratita N, Wuthiekanun V, Limmathurotsakul D, Thanwisai A, Chantratita W, Day NP, Peacock SJ. 2007. Prospective clinical evaluation of the accuracy of 16S rRNA real-time PCR assay for the diagnosis of melioidosis. Am J Trop Med Hyg 77:814–817. 139. Richardson LJ, Kaestli M, Mayo M, Bowers JR, Tuanyok A, Schupp J, Engelthaler D, Wagner DM, Keim PS, Currie BJ. 2012. Towards a rapid molecular diagnostic for melioidosis: comparison of DNA extraction methods from clinical specimens. J Microbiol Methods 88:179– 181. 140. Kaestli M, Richardson LJ, Colman RE, Tuanyok A, Price EP, Bowers JR, Mayo M, Kelley E, Seymour ML, Sarovich DS, Pearson T, Engelthaler DM, Wagner DM, Keim PS, Schupp JM, Currie BJ. 2012. Comparison of TaqMan PCR assays for detection of the melioidosis agent Burkholderia pseudomallei in clinical specimens. J Clin Microbiol 50:2059–2062. 141. Chantratita N, Meumann E, Thanwisai A, Limmathurotsakul D, Wuthiekanun V, Wannapasni S, Tumapa S, Day NP, Peacock SJ. 2008. Loop-mediated isothermal amplification method targeting the TTS1 gene cluster for detection of Burkholderia pseudomallei and diagnosis of melioidosis. J Clin Microbiol 46:568–573. 142. Gilligan PH, Gage PA, Bradshaw LM, Schidlow DV, DeCicco BT. 1985. Isolation medium for the recovery of Pseudomonas cepacia from respiratory secretions of patients with cystic fibrosis. J Clin Microbiol 22:5–8. 143. Welch DF, Muszynski MJ, Pai CH, Marcon MJ, Hribar MM, Gilligan PH, Matsen JM, Ahlin PA, Hilman BC, Chartrand SA. 1987. Selective and differential medium for recovery of Pseudomonas cepacia from the respiratory tracts of patients with cystic fibrosis. J Clin Microbiol 25:1730–1734. 144. Henry DA, Campbell ME, LiPuma JJ, Speert DP. 1997. Identification of Burkholderia cepacia isolates from patients with cystic fibrosis and use of a simple new selective medium. J Clin Microbiol 35:614–619. 145. Vermis K, Vandamme PA, Nelis HJ. 2003. Burkholderia cepacia complex genomovars: utilization of carbon sources, susceptibility to antimicrobial agents and growth on selective media. J Appl Microbiol 95:1191–1199. 146. Wright RM, Moore JE, Shaw A, Dunbar K, Dodd M, Webb K, Redmond AO, Crowe M, Murphy PG, Peacock S, Elborn JS. 2001. Improved cultural detection of Burkholderia cepacia from sputum in patients with cystic fibrosis. J Clin Pathol 54:803–805. 147. Wuthiekanun V, Dance DA, Wattanagoon Y, Supputtamongkol Y, Chaowagul W, White NJ. 1990. The use of selective media for the isolation of Pseudomonas pseudomallei in clinical practice. J Med Microbiol 33:121–126. 148. Howard K, Inglis TJ. 2003. Novel selective medium for isolation of Burkholderia pseudomallei. J Clin Microbiol 41:3312–3316. 149. Peacock SJ, Chieng G, Cheng AC, Dance DA, Amornchai P, Wongsuvan G, Teerawattanasook N, Chierakul W, Day NP, Wuthiekanun V. 2005. Comparison of Ashdown’s medium, Burkholderia cepacia medium, and Burkholderia pseudomallei selective agar for clinical isolation of Burkholderia pseudomallei. J Clin Microbiol 43:5359– 5361. 150. Denton M, Hall MJ, Todd NJ, Kerr KG, Littlewood JM. 2000. Improved isolation of Stenotrophomonas maltophilia from the sputa of patients with cystic fibrosis using a selective medium. Clin Microbiol Infect 6:397–398. 151. McMenamin JD, Zaccone TM, Coenye T, Vandamme P, LiPuma JJ. 2000. Misidentification of Burkholderia cepacia in US cystic fibrosis treatment centers: an analysis of 1,051 recent sputum isolates. Chest 117:1661–1665. 152. Brisse S, Stefani S, Verhoef J, Van Belkum A, Vandamme P, Goessens W. 2002. Comparative evaluation

153.

154.

155.

156.

157.

158.

159.

160.

161.

162.

163.

164.

165.

166.

809

of the BD Phoenix and VITEK 2 automated instruments for identification of isolates of the Burkholderia cepacia complex. J Clin Microbiol 40:1743–1748. LiPuma JJ. 1998. Burkholderia cepacia epidemiology and pathogenesis: implications for infection control. Curr Opin Pulm Med 4:337–341. LiPuma JJ, Dulaney BJ, McMenamin JD, Whitby PW, Stull TL, Coenye T, Vandamme P. 1999. Development of rRNA-based PCR assays for identification of Burkholderia cepacia complex isolates recovered from cystic fibrosis patients. J Clin Microbiol 37:3167–3170. Mahenthiralingam E, Drevinek P. 2007. Comparative genomics of Burkholderia species, p 53–79. In Coenye T, Vandamme P (ed), Burkholderia: Molecular Microbiology and Genomics. Horizon Bioscience, Wymondham, United Kingdom. Vandamme P, Henry D, Coenye T, Nzula S, Vancanneyt M, LiPuma JJ, Speert DP, Govan JR, Mahenthiralingam E. 2002. Burkholderia anthina sp. nov. and Burkholderia pyrrocinia, two additional Burkholderia cepacia complex bacteria, may confound results of new molecular diagnostic tools. FEMS Immunol Med Microbiol 33:143–149. Stead DE. 1992. Grouping of plant-pathogenic and some other Pseudomonas spp. by using cellular fatty acid profiles. Int J Syst Evol Microbiol 42:281–285. Vermis K, Coenye T, LiPuma JJ, Mahenthiralingam E, Nelis HJ, Vandamme P. 2004. Proposal to accommodate Burkholderia cepacia genomovar VI as Burkholderia dolosa sp. nov. Int J Syst Evol Microbiol 54:689–691. Whitby PW, Pope LC, Carter KB, LiPuma JJ, Stull TL. 2000. Species-specific PCR as a tool for the identification of Burkholderia gladioli. J Clin Microbiol 38:282–285. Coenye T, Mahenthiralingam E, Henry D, LiPuma JJ, Laevens S, Gillis M, Speert DP, Vandamme P. 2001. Burkholderia ambifaria sp. nov., a novel member of the Burkholderia cepacia complex including biocontrol and cystic fibrosis-related isolates. Int J Syst Evol Microbiol 51:1481–1490. Mahenthiralingam E, Bischof J, Byrne SK, Radomski C, Davies JE, Av-Gay Y, Vandamme P. 2000. DNAbased diagnostic approaches for identification of Burkholderia cepacia complex, Burkholderia vietnamiensis, Burkholderia multivorans, Burkholderia stabilis, and Burkholderia cepacia genomovars I and III. J Clin Microbiol 38:3165– 3173. Vanlaere E, Baldwin A, Gevers D, Henry D, De Brandt E, LiPuma JJ, Mahenthiralingam E, Speert DP, Dowson C, Vandamme P. 2009. Taxon K, a complex within the Burkholderia cepacia complex, comprises at least two novel species, Burkholderia contaminans sp. nov. and Burkholderia lata sp. nov. Int J Syst Evol Microbiol 59:102–111. Vanlaere E, LiPuma JJ, Baldwin A, Henry D, De Brandt E, Mahenthiralingam E, Speert D, Dowson C, Vandamme P. 2008. Burkholderia latens sp. nov., Burkholderia diffusa sp. nov., Burkholderia arboris sp. nov., Burkholderia seminalis sp. nov. and Burkholderia metallica sp. nov., novel species within the Burkholderia cepacia complex. Int J Syst Evol Microbiol 58:1580–1590. Vermis K, Coenye T, Mahenthiralingam E, Nelis HJ, Vandamme P. 2002. Evaluation of species-specific recAbased PCR tests for genomovar level identification within the Burkholderia cepacia complex. J Med Microbiol 51:937– 940. Payne GW, Vandamme P, Morgan SH, LiPuma JJ, Coenye T, Weightman AJ, Jones TH, Mahenthiralingam E. 2005. Development of a recA gene-based identification approach for the entire Burkholderia genus. Appl Environ Microbiol 71:3917–3927. Vermis K, Vandekerckhove C, Nelis HJ, Vandamme PA. 2002. Evaluation of restriction fragment length

810 n

167.

168.

169.

170.

171.

172.

173.

174.

175.

176.

177.

178.

BACTERIOLOGY

polymorphism analysis of 16S rDNA as a tool for genomovar characterisation within the Burkholderia cepacia complex. FEMS Microbiol Lett 214:1–5. Baldwin A, Mahenthiralingam E, Thickett KM, Honeybourne D, Maiden MC, Govan JR, Speert DP, Lipuma JJ, Vandamme P, Dowson CG. 2005. Multilocus sequence typing scheme for the Burkholderia cepacia complex. J Clin Microbiol 43:4665–4673. Spilker T, Baldwin A, Bumford A, Dowson CG, Mahenthiralingam E, LiPuma JJ. 2009. Expanded multilocus sequence typing for Burkholderia species. J Clin Microbiol 47:2607–2610. Brisse S, Verduin CM, Milatovic D, Fluit A, Verhoef J, Laevens S, Vandamme P, Tümmler B, Verbrugh HA, van Belkum A. 2000. Distinguishing species of the Burkholderia cepacia complex and Burkholderia gladioli by automated ribotyping. J Clin Microbiol 38:1876–1884. Coenye T, Schouls LM, Govan JR, Kersters K, Vandamme P. 1999. Identification of Burkholderia species and genomovars from cystic fibrosis patients by AFLP fingerprinting. Int J Syst Bacteriol 49:1657–1666. Vandamme P, Holmes B, Vancanneyt M, Coenye T, Hoste B, Coopman R, Revets H, Lauwers S, Gillis M, Kersters K, Govan JR. 1997. Occurrence of multiple genomovars of Burkholderia cepacia in cystic fibrosis patients and proposal of Burkholderia multivorans sp. nov. Int J Syst Bacteriol 47:1188–1200. Degand N, Carbonnelle E, Dauphin B, Beretti JL, Le Bourgeois M, Sermet-Gaudelus I, Segonds C, Berche P, Nassif X, Ferroni A. 2008. Matrix-assisted laser desorption ionization–time of flight mass spectrometry for identification of nonfermenting gram-negative bacilli isolated from cystic fibrosis patients. J Clin Microbiol 46:3361–3367. Fehlberg LC, Andrade LH, Assis DM, Pereira RH, Gales AC, Marques EA. 2013. Performance of MALDIToF MS for species identification of Burkholderia cepacia complex clinical isolates. Diagn Microbiol Infect Dis 77:126–128. Fernandez-Olmos A, Garcia-Castillo M, Morosini MI, Lamas A, Maiz L, Canton R. 2012. MALDI-TOF MS improves routine identification of non-fermenting Gram negative isolates from cystic fibrosis patients. J Cyst Fibros 11:59–62. Karger A, Stock R, Ziller M, Elschner MC, Bettin B, Melzer F, Maier T, Kostrzewa M, Scholz HC, Neubauer H, Tomaso H. 2012. Rapid identification of Burkholderia mallei and Burkholderia pseudomallei by intact cell matrixassisted laser desorption/ionisation mass spectrometric typing. BMC Microbiol 12:229. doi:10.1186/1471-218012-229. Lau SK, Tang BS, Curreem SO, Chan TM, Martelli P, Tse CW, Wu AK, Yuen KY, Woo PC. 2012. Matrixassisted laser desorption ionization–time of flight mass spectrometry for rapid identification of Burkholderia pseudomallei: importance of expanding databases with pathogens endemic to different localities. J Clin Microbiol 50:3142–3143. Marko DC, Saffert RT, Cunningham SA, Hyman J, Walsh J, Arbefeville S, Howard W, Pruessner J, Safwat N, Cockerill FR, Bossler AD, Patel R, Richter SS. 2012. Evaluation of the Bruker Biotyper and Vitek MS matrixassisted laser desorption ionization–time of flight mass spectrometry systems for identification of nonfermenting gram-negative bacilli isolated from cultures from cystic fibrosis patients. J Clin Microbiol 50:2034–2039. Miñán A, Bosch A, Lasch P, Stämmler M, Serra DO, Degrossi J, Gatti B, Vay C, D’aquino M, Yantorno O, Naumann D. 2009. Rapid identification of Burkholderia cepacia complex species including strains of the novel

179.

180.

181.

182.

183.

184.

185.

186.

187.

188.

189.

190.

191.

192.

Taxon K, recovered from cystic fibrosis patients by intact cell MALDI-ToF mass spectrometry. Analyst 134:1138– 1148. Lambiase A, Del Pezzo M, Cerbone D, Raia V, Rossano F, Catania MR. 2013. Rapid identification of Burkholderia cepacia complex species recovered from cystic fibrosis patients using matrix-assisted laser desorption ionization time-of-flight mass spectrometry. J Microbiol Methods 92:145–149. Desai AP, Stanley T, Atuan M, McKey J, LiPuma JJ, Rogers B, Jerris R. 2012. Use of matrix assisted laser desorption ionisation–time of flight mass spectrometry in a paediatric clinical laboratory for identification of bacteria commonly isolated from cystic fibrosis patients. J Clin Pathol 65:835–838. Vanlaere E, Sergeant K, Dawyndt P, Kallow W, Erhard M, Sutton H, Dare D, Devreese B, Samyn B, Vandamme P. 2008. Matrix-assisted laser desorption ionisation-timeof of-flight mass spectrometry of intact cells allows rapid identification of Burkholderia cepacia complex. J Microbiol Methods 75:279–286. Moore RA, Reckseidler-Zenteno S, Kim H, Nierman W, Yu Y, Tuanyok A, Warawa J, DeShazer D, Woods DE. 2004. Contribution of gene loss to the pathogenic evolution of Burkholderia pseudomallei and Burkholderia mallei. Infect Immun 72:4172–4187. Glass MB, Popovic T. 2005. Preliminary evaluation of the API 20NE and RapID NF Plus systems for rapid identification of Burkholderia pseudomallei and B. mallei. J Clin Microbiol 43:479–483. Inglis TJ, Aravena-Roman M, Ching S, Croft K, Wuthiekanun V, Mee BJ. 2003. Cellular fatty acid profile distinguishes Burkholderia pseudomallei from avirulent Burkholderia thailandensis. J Clin Microbiol 41:4812–4814. Amornchai P, Chierakul W, Wuthiekanun V, Mahakhunkijcharoen Y, Phetsouvanh R, Currie BJ, Newton PN, van Vinh Chau N, Wongratanacheewin S, Day NP, Peacock SJ. 2007. Accuracy of Burkholderia pseudomallei identification using the API 20NE system and a latex agglutination test. J Clin Microbiol 45:3774–3776. Deepak RN, Crawley B, Phang E. 2008. Burkholderia pseudomallei identification: a comparison between the API 20NE and VITEK2GN systems. Trans R Soc Trop Med Hyg 102(Suppl 1):S42–S4. Inglis TJ, Chiang D, Lee GS, Chor-Kiang L. 1998. Potential misidentification of Burkholderia pseudomallei by API 20NE. Pathology 30:62–64. Inglis TJ, Merritt A, Chidlow G, Aravena-Roman M, Harnett G. 2005. Comparison of diagnostic laboratory methods for identification of Burkholderia pseudomallei. J Clin Microbiol 43:2201–2206. Lowe P, Engler C, Norton R. 2002. Comparison of automated and nonautomated systems for identification of Burkholderia pseudomallei. J Clin Microbiol 40:4625–4627. Lowe P, Haswell H, Lewis K. 2006. Use of various common isolation media to evaluate the new VITEK 2 colorimetric GN card for identification of Burkholderia pseudomallei. J Clin Microbiol 44:854–856. Podin Y, Kaestli M, McMahon N, Hennessy J, Ngian HU, Wong JS, Mohana A, Wong SC, William T, Mayo M, Baird RW, Currie BJ. 2013. Reliability of automated biochemical identification of Burkholderia pseudomallei is regionally dependent. J Clin Microbiol 51:3076–3078. Goris J, De Vos P, Coenye T, Hoste B, Janssens D, Brim H, Diels L, Mergeay M, Kersters K, Vandamme P. 2001. Classification of metal-resistant bacteria from industrial biotopes as Ralstonia campinensis sp. nov., Ralstonia metallidurans sp. nov. and Ralstonia basilensis Steinle et al. 1998 emend. Int J Syst Evol Microbiol 51:1773– 1782.

43. Miscellaneous Gram-Negative Bacteria n 193. Coenye T, Vandamme P, LiPuma JJ. 2002. Infection by Ralstonia species in cystic fibrosis patients: identification of R. pickettii and R. mannitolilytica by polymerase chain reaction. Emerg Infect Dis 8:692–696. 194. Mellmann A, Cloud J, Maier T, Keckevoet U, Ramminger I, Iwen P, Dunn J, Hall G, Wilson D, Lasala P, Kostrzewa M, Harmsen D. 2008. Evaluation of matrixassisted laser desorption ionization–time-of-flight mass spectrometry in comparison to 16S rRNA gene sequencing for species identification of nonfermenting bacteria. J Clin Microbiol 46:1946–1954. 195. Coenye T, Liu L, Vandamme P, LiPuma JJ. 2001. Identification of Pandoraea species by 16S ribosomal DNAbased PCR assays. J Clin Microbiol 39:4452–4455. 196. Segonds C, Paute S, Chabanon G. 2003. Use of amplified ribosomal DNA restriction analysis for identification of Ralstonia and Pandoraea species: interest in determination of the respiratory bacterial flora in patients with cystic fibrosis. J Clin Microbiol 41:3415–3418. 197. Coenye T, LiPuma JJ. 2002. Use of the gyrB gene for the identification of Pandoraea species. FEMS Microbiol Lett 208:15–19. 198. Weyant RS, Moss CW, Weaver RE, Hollis DG, Jordan JG, Cook EC, Daneshvar MI. 1996. Identification of Unusual Pathogenic Gram-Negative Aerobic and Facultatively Anaerobic Bacteria, 2nd ed, vol 1. Williams and Wilkins, Baltimore, MD. 199. Carmody LA, Spilker T, LiPuma JJ. 2011. Reassessment of Stenotrophomonas maltophilia phenotype. J Clin Microbiol 49:1101–1103. 200. Burdge DR, Noble MA, Campbell ME, Krell VL, Speert DP. 1995. Xanthomonas maltophilia misidentified as Pseudomonas cepacia in cultures of sputum from patients with cystic fibrosis: a diagnostic pitfall with major clinical implications. Clin Infect Dis 20:445–448. 201. Whitby PW, Carter KB, Burns JL, Royall JA, LiPuma JJ, Stull TL. 2000. Identification and detection of Stenotrophomonas maltophilia by rRNA-directed PCR. J Clin Microbiol 38:4305–4309. 202. Giordano A, Magni A, Trancassini M, Varesi P, Turner R, Mancini C. 2006. Identification of respiratory isolates of Stenotrophomonas maltophilia by commercial biochemical systems and species-specific PCR. J Microbiol Methods 64:135–138. 203. Coenye T, Spilker T, Martin A, LiPuma JJ. 2002. Comparative assessment of genotyping methods for epidemiologic study of Burkholderia cepacia genomovar III. J Clin Microbiol 40:3300–3307. 204. Lessie TG, Hendrickson W, Manning BD, Devereux R. 1996. Genomic complexity and plasticity of Burkholderia cepacia. FEMS Microbiol Lett 144:117–128. 205. Inglis TJ, O’Reilly L, Foster N, Clair A, Sampson J. 2002. Comparison of rapid, automated ribotyping and DNA macrorestriction analysis of Burkholderia pseudomallei. J Clin Microbiol 40:3198–3203. 206. LiPuma JJ, Dasen SE, Nielson DW, Stern RC, Stull TL. 1990. Person-to-person transmission of Pseudomonas cepacia between patients with cystic fibrosis. Lancet 336:1094–1096. 207. LiPuma JJ, Mortensen JE, Dasen SE, Edlind TD, Schidlow DV, Burns JL, Stull TL. 1988. Ribotype analysis of Pseudomonas cepacia from cystic fibrosis treatment centers. J Pediatr 113:859–862. 208. Krzewinski JW, Nguyen CD, Foster JM, Burns JL. 2001. Use of random amplified polymorphic DNA PCR to examine epidemiology of Stenotrophomonas maltophilia and Achromobacter (Alcaligenes) xylosoxidans from patients with cystic fibrosis. J Clin Microbiol 39:3597–3602. 209. Mahenthiralingam E, Campbell ME, Henry DA, Speert DP. 1996. Epidemiology of Burkholderia cepacia infection

210.

211.

212.

213.

214.

215.

216.

217.

218.

219.

220.

221.

222.

223.

224.

811

in patients with cystic fibrosis: analysis by randomly amplified polymorphic DNA fingerprinting. J Clin Microbiol 34:2914–2920. Cheng AC, Godoy D, Mayo M, Gal D, Spratt BG, Currie BJ. 2004. Isolates of Burkholderia pseudomallei from northern Australia are distinct by multilocus sequence typing, but strain types do not correlate with clinical presentation. J Clin Microbiol 42:5477–5483. Godoy D, Randle G, Simpson AJ, Aanensen DM, Pitt TL, Kinoshita R, Spratt BG. 2003. Multilocus sequence typing and evolutionary relationships among the causative agents of melioidosis and glanders, Burkholderia pseudomallei and Burkholderia mallei. J Clin Microbiol 41:2068– 2079. Currie BJ, Gal D, Mayo M, Ward L, Godoy D, Spratt BG, LiPuma JJ. 2007. Using BOX-PCR to exclude a clonal outbreak of melioidosis. BMC Infect Dis 7:68. doi:10.1186/1471-2334-7-68. Currie BJ, Haslem A, Pearson T, Hornstra H, Leadem B, Mayo M, Gal D, Ward L, Godoy D, Spratt BG, Keim P. 2009. Identification of melioidosis outbreak by multilocus variable number tandem repeat analysis. Emerg Infect Dis 15:169–174. Wuthiekanun V, Langa S, Swaddiwudhipong W, Jedsadapanpong W, Kaengnet Y, Chierakul W, Day NP, Peacock SJ. 2006. Short report: Melioidosis in Myanmar: forgotten but not gone? Am J Trop Med Hyg 75:945–946. Peacock SJ, Schweizer HP, Dance DA, Smith TL, Gee JE, Wuthiekanun V, DeShazer D, Steinmetz I, Tan P, Currie BJ. 2008. Management of accidental laboratory exposure to Burkholderia pseudomallei and B. mallei. Emerg Infect Dis 14:e2. doi:10.3201/eid1407.071501. Cheng AC, O’Brien M, Freeman K, Lum G, Currie BJ. 2006. Indirect hemagglutination assay in patients with melioidosis in northern Australia. Am J Trop Med Hyg 74:330–334. Cheng AC, Peacock SJ, Limmathurotsakul D, Wongsuvan G, Chierakul W, Amornchai P, Getchalarat N, Chaowagul W, White NJ, Day NP, Wuthiekanun V. 2006. Prospective evaluation of a rapid immunochromogenic cassette test for the diagnosis of melioidosis in northeast Thailand. Trans R Soc Trop Med Hyg 100:64–67. O’Brien M, Freeman K, Lum G, Cheng AC, Jacups SP, Currie BJ. 2004. Further evaluation of a rapid diagnostic test for melioidosis in an area of endemicity. J Clin Microbiol 42:2239–2240. Wuthiekanun V, Amornchai P, Chierakul W, Cheng AC, White NJ, Peacock SJ, Day NP. 2004. Evaluation of immunoglobulin M (IgM) and IgG rapid cassette test kits for diagnosis of melioidosis in an area of endemicity. J Clin Microbiol 42:3435–3437. Hancock RE. 1998. Resistance mechanisms in Pseudomonas aeruginosa and other nonfermentative gram-negative bacteria. Clin Infect Dis 27(Suppl 1):S93–S99. Poole K, Srikumar R. 2001. Multidrug efflux in Pseudomonas aeruginosa: components, mechanisms and clinical significance. Curr Top Med Chem 1:59–71. Aaron SD, Ferris W, Henry DA, Speert DP, MacDonald NE. 2000. Multiple combination bactericidal antibiotic testing for patients with cystic fibrosis infected with Burkholderia cepacia. Am J Respir Crit Care Med 161:1206– 1212. Hoban DJ, Bouchillon SK, Johnson JL, Zhanel GG, Butler DL, Miller LA, Poupard JA, Gemifloxacin Surveillance Study Research Group. 2001. Comparative in vitro activity of gemifloxacin, ciprofloxacin, levofloxacin and ofloxacin in a North American surveillance study. Diagn Microbiol Infect Dis 40:51–57. Milatovic D, Schmitz FJ, Verhoef J, Fluit AC. 2003. Activities of the glycylcycline tigecycline (GAR-936)

812 n

225.

226.

227.

228.

229.

230.

231.

232.

233.

234.

235.

BACTERIOLOGY

against 1,924 recent European clinical bacterial isolates. Antimicrob Agents Chemother 47:400–404. Clinical and Laboratory Standards Institute. 2005. Performance Standards for Antimicrobial Susceptibility Testing; 15th Informational Supplement. CLSI Document MI00S15. Clinical and Laboratory Standards Institute, Wayne, PA. Clinical and Laboratory Standards Institute. 2010. Methods for Antimicrobial Dilution and Disk Susceptibility Testing of Infrequently Isolated or Fastidious Bacteria; Approved Guideline—2nd Edition. CLSI Document M45-A2. Clinical and Laboratory Standards Institute, Wayne, PA. Jenney AW, Lum G, Fisher DA, Currie BJ. 2001. Antibiotic susceptibility of Burkholderia pseudomallei from tropical northern Australia and implications for therapy of melioidosis. Int J Antimicrob Agents 17:109–113. Piliouras P, Ulett GC, Ashhurst-Smith C, Hirst RG, Norton RE. 2002. A comparison of antibiotic susceptibility testing methods for cotrimoxazole with Burkholderia pseudomallei. Int J Antimicrob Agents 19:427–429. Lumbiganon P, Tattawasatra U, Chetchotisakd P, Wongratanacheewin S, Thinkhamrop B. 2000. Comparison between the antimicrobial susceptibility of Burkholderia pseudomallei to trimethoprim-sulfamethoxazole by standard disk diffusion method and by minimal inhibitory concentration determination. J Med Assoc Thai 83:856–860. Wuthiekanun V, Cheng AC, Chierakul W, Amornchai P, Limmathurotsakul D, Chaowagul W, Simpson AJ, Short JM, Wongsuvan G, Maharjan B, White NJ, Peacock SJ. 2005. Trimethoprim/sulfamethoxazole resistance in clinical isolates of Burkholderia pseudomallei. J Antimicrob Chemother 55:1029–1031. Chaowagul W, Suputtamongkul Y, Smith MD, White NJ. 1997. Oral fluoroquinolones for maintenance treatment of melioidosis. Trans R Soc Trop Med Hyg 91:599– 601. Feterl M, Govan B, Engler C, Norton R, Ketheesan N. 2006. Activity of tigecycline in the treatment of acute Burkholderia pseudomallei infection in a murine model. Int J Antimicrob Agents 28:460–464. Thamlikitkul V, Trakulsomboon S. 2006. In vitro activity of tigecycline against Burkholderia pseudomallei and Burkholderia thailandensis. Antimicrob Agents Chemother 50:1555–1557. Chaowagul W, Chierakul W, Simpson AJ, Short JM, Stepniewska K, Maharjan B, Rajchanuvong A, Busarawong D, Limmathurotsakul D, Cheng AC, Wuthiekanun V, Newton PN, White NJ, Day NP, Peacock SJ. 2005. Open-label randomized trial of oral trimethoprimsulfamethoxazole, doxycycline, and chloramphenicol compared with trimethoprim-sulfamethoxazole and doxycycline for maintenance therapy of melioidosis. Antimicrob Agents Chemother 49:4020–4025. Chierakul W, Anunnatsiri S, Short JM, Maharjan B, Mootsikapun P, Simpson AJ, Limmathurotsakul D, Cheng AC, Stepniewska K, Newton PN, Chaowagul W, White NJ, Peacock SJ, Day NP, Chetchotisakd P. 2005. Two randomized controlled trials of ceftazidime alone versus ceftazidime in combination with trimetho-

236.

237.

238.

239.

240.

241.

242.

243.

244.

245.

prim-sulfamethoxazole for the treatment of severe melioidosis. Clin Infect Dis 41:1105–1113. Chetchotisakd P, Chierakul W, Chaowagul W, Anunnatsiri S, Phimda K, Mootsikapun P, Chaisuksant S, Pilaikul J, Thinkhamrop B, Phiphitaporn S, Susaengrat W, Toondee C, Wongrattanacheewin S, Wuthiekanun V, Chantratita N, Thaipadungpanit J, Day NP, Limmathurotsakul D, Peacock SJ. 2014. Trimethoprim-sulfamethoxazole versus trimethoprim-sulfamethoxazole plus doxycycline as oral eradicative treatment for melioidosis (MERTH): a multicentre, double-blind, non-inferiority, randomised controlled trial. Lancet 383:807–814. Cheng AC, Chierakul W, Chaowagul W, Chetchotisakd P, Limmathurotsakul D, Dance DA, Peacock SJ, Currie BJ. 2008. Consensus guidelines for dosing of amoxicillinclavulanate in melioidosis. Am J Trop Med Hyg 78:208– 209. Cheng AC, Fisher DA, Anstey NM, Stephens DP, Jacups SP, Currie BJ. 2004. Outcomes of patients with melioidosis treated with meropenem. Antimicrob Agents Chemother 48:1763–1765. Cheng AC, Limmathurotsakul D, Chierakul W, Getchalarat N, Wuthiekanun V, Stephens DP, Day NP, White NJ, Chaowagul W, Currie BJ, Peacock SJ. 2007. A randomized controlled trial of granulocyte colony-stimulating factor for the treatment of severe sepsis due to melioidosis in Thailand. Clin Infect Dis 45:308–314. Cheng AC, Stephens DP, Currie BJ. 2004. Granulocytecolony stimulating factor (G-CSF) as an adjunct to antibiotics in the treatment of pneumonia in adults. Cochrane Database Syst Rev 2004:CD004400. doi:10.1002/ 14651858.CD004400.pub2. Heine HS, England MJ, Waag DM, Byrne WR. 2001. In vitro antibiotic susceptibilities of Burkholderia mallei (causative agent of glanders) determined by broth microdilution and E-test. Antimicrob Agents Chemother 45: 2119–2121. Lipsitz R, Garges S, Aurigemma R, Baccam P, Blaney DD, Cheng AC, Currie BJ, Dance D, Gee JE, Larsen J, Limmathurotsakul D, Morrow MG, Norton R, O’Mara E, Peacock SJ, Pesik N, Rogers LP, Schweizer HP, Steinmetz I, Tan G, Tan P, Wiersinga WJ, Wuthiekanun V, Smith TL. 2012. Workshop on treatment of and postexposure prophylaxis for Burkholderia pseudomallei and B. mallei infection, 2010. Emerg Infect Dis 18:e2. doi:10.3201/eid1812.120638. Garrison MW, Anderson DE, Campbell DM, Carroll KC, Malone CL, Anderson JD, Hollis RJ, Pfaller MA. 1996. Stenotrophomonas maltophilia: emergence of multidrug-resistant strains during therapy and in an in vitro pharmacodynamic chamber model. Antimicrob Agents Chemother 40:2859–2864. San Gabriel P, Zhou J, Tabibi S, Chen Y, Trauzzi M, Saiman L. 2004. Antimicrobial susceptibility and synergy studies of Stenotrophomonas maltophilia isolates from patients with cystic fibrosis. Antimicrob Agents Chemother 48:168–171. Barbaro DJ, Mackowiak PA, Barth SS, Southern PM Jr. 1987. Pseudomonas testosteroni infections: eighteen recent cases and a review of the literature. Rev Infect Dis 9:124– 129.

Acinetobacter, Chryseobacterium, Moraxella, and Other Nonfermentative Gram-Negative Rods* MARIO VANEECHOUTTE, ALEXANDR NEMEC, PETER KÄMPFER, PIET COOLS, AND GEORGES WAUTERS

44 their clinical importance is mostly restricted to that of opportunistic pathogens, except, e.g., for Elizabethkingia meningoseptica, Moraxella lacunata (eye infections), M. catarrhalis (respiratory tract infections), and species of the Acinetobacter calcoaceticus-Acinetobacter baumannii complex. A. baumannii ventilator-associated pneumonia and bloodstream infections have been documented to be associated with a high degree of mortality and morbidity (4). Particular manifestations of A. baumannii are its implication in severely war-wounded soldiers (5), from which stems its popular designation “Iraqibacter,” and in victims of natural disasters (6). The clinical impact of infections with A. baumannii is a continuous source of debate (7, 8). Indeed, although severe infections with A. baumannii have been documented, colonization is much more frequent than infection, and differentiation between these conditions can be difficult. Still, although uncommon, community-acquired infections with A. baumannii occur. In particular, community-acquired pneumonia with A. baumannii is increasingly reported from tropical areas, like Southeast Asia and tropical Australia (9, 10). The clinical role of the closely related species Acinetobacter pittii and Acinetobacter nosocomialis resembles that of A. baumannii, although, compared to these two species, A. baumannii is more often associated with multidrug resistance and epidemic spread in hospitals and possibly also with higher mortality among patients with systemic infections (11, 12). Other Acinetobacter species occasionally associated with human infections are listed in Table 1. A. johnsonii, A. lwoffii, and A. radioresistens seem to be common inhabitants of human skin (13). A. lwoffii was a frequent species in clinical specimens during an 8-year study in a university hospital, where it was isolated mainly from blood or intravascular lines (14). Several species, A. ursingii in particular, have been found to cause bloodstream infections in hospitalized patients (15–18), while A. junii and A. soli have also been implicated in outbreaks of neonatal infections (19, 20). A. parvus is regularly isolated from blood cultures (18, 21), but is often overlooked because of its small colonies and misidentification by API 20NE as A. lwoffii (our unpublished data). Many of the infections with these species are related to intravascular catheters or have another iatrogenic

TAXONOMY The organisms covered in this chapter belong to a group of taxonomically and phylogenetically diverse, Gram-negative, nonfermentative rods and coccobacilli. Still, several of the genera dealt with belong to the same family; i.e., Acinetobacter, Moraxella, Oligella, and Psychrobacter belong to the family Moraxellaceae (Gammaproteobacteria) (1), and Balneatrix, Bergeyella, Chryseobacterium, Elizabethkingia, Empedobacter, Myroides, Sphingobacterium, Wautersiella, and Weeksella belong to the family Flavobacteriaceae (Bacteroidetes) (2).

DESCRIPTION OF THE AGENTS The species dealt with in this chapter all share the common phenotypic features of being catalase positive and failing to acidify the butt of Kligler iron agar (KIA) or triple sugar iron (TSI) agar or of oxidative-fermentative media, indicating their inability to metabolize carbohydrates by the fermentative pathway. These organisms grow significantly better under aerobic than under anaerobic conditions, and many, i.e., those species that can use only oxygen as the final electron acceptor in the respiratory pathway, fail to grow anaerobically at all.

EPIDEMIOLOGY AND TRANSMISSION Most of the organisms described in this chapter are found in the environment, i.e., soil and water. For methylobacteria, tap water has been implicated as a possible agent of transmission in hospital environments, and methods for monitoring water systems for methylobacteria have been described previously (3). No person-to-person spread has been documented for the species covered in this chapter, except for Acinetobacter and Moraxella catarrhalis.

CLINICAL SIGNIFICANCE Although for almost all of the species in this chapter, case reports, e.g., of meningitis and endocarditis, can be found, *This chapter contains information presented by Mario Vaneechoutte, Lenie Dijkshoorn, Alexandr Nemec, Peter Kämpfer, and Georges Wauters in chapter 42 of the 10th edition of this Manual.

doi:10.1128/9781555817381.ch44

813

814 n BACTERIOLOGY

100 94 88 0 100 100 100 0 100 65 94 100 18 65 0 0

100 100 /g 100 0 100 100 0 /g 0 100 100 78 100 100 100

A. ursingii (30)

63 100 100 0 100 63 100 0 100 63 94 100 0 25 0 0

A. soli (8)

0 0 100 0 100 0 100 0 0 0 100 0 100 0 13 100

A. schindleri (22)

100 90 100 100 100 100 100 0 90 0 100 100 95 75 0 0

A. radioresistens (12)

95 85 100 100 100 95 100 0 95 0 100 100 20 85 0 0

100 91 100 100 100 100 100 0 91 0 100 100 100 100 0 0

A. parvus (10)

88 100 100 100 100 88 100 0 96 0 96 100 88 84 0 0

0 0 0 0 0e 100 0 0

0 94 100 82

0 87 100 0

0 0 0 7 25f 100 0 7

0 0 90 0

0 100 100 0

0 100 100 0

0 88 100 100

0 0 100 0

0 0 100 88 18 0 82 0 0 0 100 0 0 0 88 100

0 0 87 93 27 0 80 0 0 0 93 93 0 0 0 47

0 79 0 0 75 79 70 0 75 0 0 100 85 14 0 0 0 0 0 0 0 0 100 93 50 7 0 71 0 0 25f 0

0 0 0 0 0 0 0 0 0 0 0 0 0 0 0 0

92 0 100 83 0 100 0 0 100 0 0 100 100 100 0 0

38 0 0 0 0 64 59 0 95 0 0 100 0 0 0 0

100 100 100 100 100 100 100 100 88 0 100 100 100 100 0 75

100 0 0 0 97f 100 97 0 97 0 0 100 0 0 0 0

A. lwoffii (14)

0 0 100e 88

A. johnsonii (20)

0 0 100 0

A. junii (15)

A. bereziniae (16)

10 100 100 95

A. haemolyticus (17)

A. beijerinckii (16)

95 100 100 100

A. gyllenbergii (9)

A. pittii (20)c

100d 0 100 9 100 100 100 91

A. guillouiae (17)

A. nosocomialis (20)c

Growth at: 44°C 41°C 37°C Acidification of D-glucose Assimilation of: Adipate β-Alanine 4-Aminobutyrate L-Arginine L-Aspartate Azelate Citrate (Simmons) D-Glucose Glutarate Histamine L-Histidine DL-Lactate Malonate Phenylacetate Gelatinase Hemolysis of sheep blood

A. baumannii (25)b,c

Characteristic

A. calcoaceticus (11)c

TABLE 1 Oxidase-negative, indole-negative, nonfermentative, Gram-negative rods: the genus Acinetobactera

a

All data were provided by one of the authors (A. Nemec) using standardized tests described in detail previously (67). Numbers in parentheses after organism names are numbers of strains tested. Species of the A. calcoaceticus-A. baumannii complex (65). d The numbers are percentages of strains positive in a given test. Carbon assimilation tests were evaluated after 6 days of incubation at 30°C. e Growth tested at 38 instead of 37°C. f Weak growth of most strains with positive reactions. g Unreproducible or delayed growth of most strains. b c

origin (15, 22), and their course is generally benign. For various other named or yet-unnamed Acinetobacter species, although recovered from clinical specimens (23, 24), a possible role in infection has not been documented. Moraxella species are rare agents of infections (conjunctivitis, keratitis, meningitis, septicemia, endocarditis, arthritis, and otolaryngologic infections) (25–27), but M. catarrhalis has been reported to cause sinusitis and otitis media by contiguous spread of the organisms from a colonizing focus in the respiratory tract (25). However, isolation of M. catarrhalis from the upper respiratory tract (i.e., a throat culture) of children with otitis media or sinusitis does not provide evidence that the isolate is the cause of these infections, because M. catarrhalis is present frequently as a commensal of the upper respiratory tract in children (28). Isolates from sinus aspirates and middle ear specimens obtained by tympanocentesis should be identified and reported. Similarly, little is known about the pathogenesis of lower respiratory tract infection in adults with chronic lung diseases, although a clear pathogenic role may be assigned to this species because M. catarrhalis is not a frequent commensal of the upper respiratory tract in adults (28) and because examination of Gram-stained smears of sputum specimens from patients with exacerbations of bronchitis and pneumonia due to M. catarrhalis usually reveals an abundance of leukocytes, the presence of many Gram-negative diplococci as the exclusive or predominant bacterial cell type, and the

presence of intracellular Gram-negative diplococci. Such specimens may yield M. catarrhalis in virtually pure culture, and the organism should be identified and reported. Furthermore, M. nonliquefaciens (29, 30) and M. osloensis (31, 32) are the two species most frequently isolated, approximately in equal numbers, from nonrespiratory clinical material, especially blood cultures from patients at risk. M. canis has been isolated from dog bite wounds (33) and from debilitated patients (27). M. lacunata has been involved in eye infections and in infective endocarditis (34, 35).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS Standard methods for collection, transport, and storage of specimens as detailed in chapters 11 and 18 are satisfactory for this group of organisms. The only fastidious species handled in this chapter are Asaia species, Granulibacter bethesdensis, Methylobacterium species, and some Moraxella species; these should be cultured on media containing blood.

DIRECT EXAMINATION There are no characteristics available that can help to recognize the species dealt with in this chapter by means of direct microscopic examination of the samples. On Gram stain, organisms appear as Gram-negative rods, coccobacilli, or

44. Nonfermentative Gram-Negative Rods n 815

diplococci. Neither direct antigen tests nor molecular genetic tests to use directly on clinical materials have been developed.

ISOLATION PROCEDURES Initial incubation should be at 35 to 37°C, although some strains, among them many of the pink-pigmented species, grow better at or below 30°C and may be detected only on plates left at room temperature. In such cases, all tests should be carried out at room temperature. In fact, some of the commercial kits, such as the API 20NE, are designed to be incubated at 30°C. Growth on certain selective primary media, e.g., MacConkey agar, is variable and may be influenced by lotto-lot variations in the composition of media. Gramnegative, nonfermentative bacteria (GNF) that grow on MacConkey agar generally form colorless colonies, although some form lavender or purple colonies due to uptake of crystal violet contained in the agar medium. Selective media have been described for Acinetobacter spp. (36) and for Moraxella spp. (37).

IDENTIFICATION This chapter provides an overview in Fig. 1, which provides a key to the five large groups that can be distinguished among the species described in this chapter. This key is

based only on colony color (pink or not) and the presence or absence of oxidase, of benzyl arginine aminopeptidase (trypsin) activity, and of the production of indole. Figure 1 refers to Tables 1 and 3 to 6, which provide further keys to identify the species of these five groups on the basis of biochemical reactions. Results for enzymatic reactions can be read within hours or up to 2 days of incubation, whereas results of carbon source assimilation tests (Acinetobacter) and acid production from carbohydrates are read after up to 6 and 7 days, respectively. For each group of closely related species, we present their taxonomic history (explaining the use of other names in the past and the taxonomic changes introduced since the previous edition), address the clinical importance of the species, and describe the phenotypic data that are useful to differentiate this group from other groups and to differentiate the species within this group. When relevant, antibiotic susceptibility characteristics and treatment options are discussed immediately; otherwise, they are discussed at the end of each section for the five large groups in this chapter. Although several of the genera discussed in this chapter comprise many more species than the ones addressed here, we focus on those species that can be isolated from clinical samples.

Classical Biochemical Identification Schemes Presented in This Chapter For all the species in this chapter, except those of the genus Acinetobacter, the biochemical tests listed have been carried

FIGURE 1 Identification of miscellaneous GNF. The organisms covered in this chapter belong to a group of taxonomically diverse, Gram-negative, nonfermentative rods and coccobacilli. They all share the common phenotypic features of failing to acidify the butt of KIA or TSI agar or of oxidative-fermentative media, indicating their inability to metabolize carbohydrates by the fermentative pathway. These characteristics are shared with those of the species of the emended genus Pseudomonas (chapter 42) and those of the species of genera that previously were named as Pseudomonas (chapter 43). a, G. bethesdensis grows slowly and poorly on SBA. A. parvus forms small colonies as well, but these are already visible after 24 h of incubation. doi:10.1128/9781555817381.ch44.f1

816 n BACTERIOLOGY

out by one of us (G. Wauters), according to standardized protocols, described in detail in chapter 33. The limited number of tests that have been used to discriminate between the species dealt with in this chapter have been selected because they can be carried out easily and quickly, because they mostly yield uniform results per group or species, and because they are highly discriminatory. For the genus Acinetobacter, data based on standardized physiological and nutritional tests were provided by one of the authors (A. Nemec) (see footnotes to Table 1).

Automated, Commercially Available Phenotypic Identification Systems Traditional diagnostic systems, e.g., those based on oxidation-fermentation media, aerobic low-peptone media, or buffered single substrates, have now been replaced in many laboratories by commercial kits or automated systems like the VITEK 2 (bioMérieux, Marcy l’Etoile, France) and the Phoenix (BD Diagnostic Systems, Sparks, MD). The ability of commercial kits to identify this group of GNF is variable and often results in identification to the genus or group level only, necessitating the use of supplemental biochemical testing for species identification. O’Hara and Miller (38), using the VITEK 2 ID-GNB identification card, reported that of 103 glucose-fermenting and nonfermenting, nonenteric strains, 88 (85.4%) were correctly identified at probability levels ranging from excellent to good and that 10 (9.7%) were correctly identified at a low level of discrimination, for a total of 95.1% accuracy within this group. Bosshard et al. (39) compared 16S rRNA gene sequencing for the identification of clinically relevant isolates of GNF (nonPseudomonas aeruginosa) with two commercially available identification systems (API 20NE and VITEK 2 fluorescent card; bioMérieux). By 16S rRNA gene sequence analysis, 92% of the isolates were assigned to species level and 8% to genus level. Using API 20NE, 54% of the isolates were assigned to species level, 7% were assigned to genus level, and 39% of the isolates could not be discriminated at any taxonomic level. The respective numbers for VITEK 2 were 53, 1, and 46%. Fifteen percent and 43% of the isolates corresponded to species not included in the API 20NE and VITEK 2 databases, respectively. Altogether, commercial identification systems can be useful for identification of organisms commonly found in clinical specimens, like Enterobacteriaceae. However, for rare organisms the performance of these systems can be poor.

Chemotaxonomic Methods The fatty acid profiles for the most common species of GNF have been published (40).

Matrix-Assisted Laser Desorption Ionization–Timeof-Flight Mass Spectrometry Few developments in clinical bacteriology have had as rapid and profound an impact on identification of microorganisms as matrix-assisted laser desorption ionization–time-of-flight mass spectrometry (MALDI-TOF MS) (41, 42). Conventional identification methods rely on biochemical properties and require sometimes lengthy incubation procedures, whereas MALDI-TOF MS can identify bacteria within minutes directly from colonies grown on culture plates. Two commercial platforms, Bruker Biotyper (Bruker Daltonics, Billerica, MA) and VITEK MS (bioMérieux), are available,

and although the initial cost of mass spectrometry equipment and maintenance costs are relatively high, the additional identification costs per isolate are minimal. Studies on the reliability and accuracy of MALDI-TOF MS in identifying species dealt with in this chapter are summarized in Table 2. MALDI-TOF MS appears to be a promising tool for identification of these species, with a reported 80.4% correct species identifications and 92.3% correct genus identifications (43, 44), with most difficulties in obtaining a correct identification due to a missing spectrum in the database at that time. These problems can be overcome by creating new databases or by adding missing spectra (43, 44). The accuracy of both MALDI-TOF MS systems was found to be similar for identifying GNF (45), and both systems demonstrated performance superior to that of conventional methods (45). Mellmann and coworkers (46) reported 98.75% interlaboratory reproducibility among eight laboratories in a study of MALDI-TOF MS for identifying 480 GNF isolates. Sedo et al. (47) successfully applied MALDI-TOF MS for identification of selected Acinetobacter species, but could not differentiate between different strains belonging to the same species. Besides bacterial identification, MALDI-TOF MS is also being explored for the phenotypic detection of certain antibiotic resistance mechanisms, e.g., the detection of carbapenemase enzymatic activity in A. baumannii strains (48), and for typing. Schaller and coworkers (49) found MALDI-TOF MS to be a rapid and robust tool for typing the two subpopulations and three 16S rRNA types of M. catarrhalis described previously. Mencacci and coworkers (50) explored the potential utility of MALDI-TOF MS to detect nosocomial spread of multidrug-resistant A. baumannii outbreaks in comparison with the repetitive sequence-based PCR DiversiLab system (bioMérieux) and suggested that it shows promise in routine clinical microbiology.

DNA Sequence-Based Methods Sequence-based methods involving rRNA (16S, 16S-23S spacer, or 23S) and housekeeping genes, such as those encoding RNA polymerase subunit B (rpoB), gyrase subunit B (gyrB), or the recombination A protein (recA), have become standard techniques to identify bacteria in general (51) and have contributed to the better delineation of several of these groups and the discovery and description of new species. Because these are generally applicable methods, their application for species of this chapter is not outlined in detail. Other sequence-based methods, based on DNA array hybridization, have been used for some species of these groups (52). DNA sequence-based fingerprinting methods like amplified ribosomal DNA (rRNA gene) restriction analysis (ARDRA) (53, 54) and tDNA PCR (55, 56) have been applied for the identification of species of several groups as well. These fingerprinting approaches are also generally applicable, but they require reference fingerprint libraries and are often poorly exchangeable between different electrophoresis platforms and laboratories.

IDENTIFICATION OF THE FIVE GENOTYPIC GROUPS Oxidase-Negative GNF Acinetobacter Members of the genus Acinetobacter are strictly aerobic, oxidase-negative, catalase-positive, coccobacillary bacteria.

44. Nonfermentative Gram-Negative Rods n 817 TABLE 2

Accuracy of MALDI-TOF MS compared to other identification methodsa Species

MALDI-TOF MS result

Acinetobacter baumannii (141)b,c,d,e,f A. beijerinckii (3)e,j A. berezinae (18)e,j A. calcoaceticus (11)b,d,e

A. baumannii (135),g,h Acinetobacter sp. (6)g,i A. beijerinckii (2), A. tjernbergiae (1) A. berezinae (11),k A. genomic species 3 (7) A. calcoaceticus, Acinetobacter sp. (8),g A. genomic species 3 (2) A. genomic species 3 (8) Acinetobacter sp., A.baumannii (2) A. genomic species 3, A. pittiil Acinetobacter sp. A. guillouiae (11) A. haemolyticus (3) A. johnsonii (2),g,h A. junii,g Acinetobacter sp.g,h A. junii (7)g,h A. lwoffii (6),g,h,iAcinetobacter sp.,g Ochrobactrum anthropig A. nosocomialis (18)m A. pittii (16) A. radioresistens (3),g,h misidentificationn A. schindlerig,h A. baylyi A. ursingii,g,h Acinetobacter sp.g,h A. junii/A. haemolyticusg No identificationg,o C. indologenes (5),g no identification (1)k,o C. indologenes (1)g E. meningoseptica (2),g,h E. miricola (1)k M. catarrhalis (18)g O. anthropi (6),g,k Ochrobactrum sp. (21),g,k O. tritici (2)g,k O. intermedium (9)g,k Ochrobactrum anthropig Oligella urethralisg No identificationg R. radiobacter (3)g,h Misidentificationg,h No identificationg,h S. spiritivorumg Acinetobacter sp. (2),k Wohlfahrtiimonas chitiniclastica (3)g

A. genomic species 3 (8)e,j A. genomic species 13TU (3)e,j A. genomic species 14BJ (2)d,e,j A. genomic species 17d,j A. guillouiae (11)e A. haemolyticus (3)d A. johnsonii (5)b,f A. junii (7)d,e,f A. lwoffii (8)c,d,f A. nosocomialis (18)d A. pittii (16)d A. radioresistens (4)b,d,f,j A. schindlerif A. solie,j A. ursingii (2)f Acinetobacter sp.b Chryseobacterium gleumb C. indologenes (6)b,f Chryseobacterium spp. (1)b Elizabethkingia meningoseptica (3)b,f Moraxella catarrhalis (18)b Ochrobactrum anthropi (29)b,f O. intermedium (9)b,f Ochrobactrum spp.b Oligella spp.b Psychrobacter phenylpyruvicusf,j Rhizobium radiobacter (3)b,f Roseomonas mucosaf,j Shewanella algaef,j Sphingobacterium spiritivorumb No identification (5)b,f

Reference(s) 42, 47, 47, 42,

126, 201–203 201 201 201, 202

201 201 42, 201 42 47 42, 47 126, 202–203 42, 126, 201 42, 126 42 42 42, 126, 202 126 201 126 202 203 44, 45, 203 203 44, 45, 126 202 203; Wauters (unpublished) Wauters (unpublished) 203 203 126 126, 203 126 126 203 44; Wauters (unpublished)

a All identifications obtained with the Bruker MALDI-TOF MS system, except for those of reference 45, which compared the Bruker and VITEK systems. Numbers in parentheses after organism names are numbers of strains tested. b Identification by conventional methods, with discrepant results resolved by 16S rRNA gene sequencing. c Conventional biochemical identification. d Identification by ARDRA (61), rRNA intergenic spacer (ITS), recA sequencing, and/or blaOXA-51 PCR. e Identification by blaOXA-51 PCR and/or rpoB gene sequencing. f Identification by 16S rRNA gene sequencing. g Direct transfer method. h Identification also taking into account the rule of thumb that the difference between the first two species listed is larger than 0.2 if the first log score is 1 mm in diameter after 24 h of incubation. Colonies of M. catarrhalis grow well on both blood and chocolate agars, and some strains also grow well on modified ThayerMartin and other selective media. Colonies are generally gray to white, opaque, and smooth and measure about 1 to 3 mm after 24 h of incubation. Characteristically, the colonies may be nudged intact across the plate with a bacteriological loop like a “hockey puck” and can be removed from the agar entirely, being very consistent. Most M. canis colonies resemble those of the Enterobacteriaceae (large, smooth colonies) and may produce a brown pigment when grown on starch-containing Mueller-Hinton agar (33). Some strains may also produce very slimy colonies resembling colonies of Klebsiella pneumoniae (33). M. nonliquefaciens forms smooth, translucent to semiopaque colonies 0.1 to 0.5 mm in diameter after 24 h and 1 mm in diameter after 48 h of growth on SBA plates. Occasionally, these colonies spread and pit the agar. The colonial morphologies of M. lincolnii (103), M. osloensis, and P. phenylpyruvicus (formerly M. phenylpyruvica) are similar, but pitting is rare. On the other hand, pitting is common with M. lacunata, whose colonies are smaller and form dark haloes on chocolate agar. Rod-shaped Moraxella species, especially M. atlantae and M. lincolnii, are more fastidious and display smaller colonies on SBA, 104 for F. novicida (9, 22). Levels of virulence also differ between F. tularensis subspecies; F. tularensis subsp. tularensis displays an LD100 of 40 whole-genome sequences have been completed for Francisella species, including F. tularensis subsp. tularensis, F. tularensis subsp. holarctica, F. tularensis subsp. mediasiatica, F. novicida, F. philomiragia, F. hispanensis, F. noatunensis, and Wolbachia persica (3). The size of Francisella genomes is ∼1.9 Mb, with a G+C content of ∼32% (http://www.ncbi.nlm.nih.gov/genome/?term= Francisella). All Franicsella genomes are comprised of a single chromosome. Plasmids are unique to some environmental Francisella spp., including F. novicida and F. philomiragia, and are not conserved across all members of a given species (http://www.ncbi.nlm.nih.gov/genome/?term=Fran cisella). Whole-genome analyses indicate that F. tularensis, the causative agent of tularemia, contains only seven unique genes in comparison to other members of the Franicsella genus (3). These genes encode proteins predicted to be involved in exopolysaccharide and lipopolysaccharide synthesis as well as a hypothetical membrane protein (3).

EPIDEMIOLOGY AND TRANSMISSION F. tularensis has been recovered only from the Northern Hemisphere, with the distribution of the subspecies varying throughout this region (1, 2, 25) (Table 3). Infections caused by F. tularensis subsp. tularensis occur only in North America, whereas F. tularensis subsp. holarctica has a much wider distribution, causing disease throughout the Northern Hemi-

TABLE 1 Differentiation of Francisella from similar Gram-negative generaa Test Oxidase Urease Gram stain morphology Specimen source

X and/or V factor requirement Cysteine enhancement Motility Major CFAe

Pasteurella multocida

F. tularensis

Brucella spp.

Bartonella spp.b

Acinetobacter spp. Haemophilus spp.c Yersinia pestis

− − Very tiny ccb Ulcer, wound, blood, lymph node aspirates, respiratory −

+ + Tiny ccb

− − Thin rod

− − Broad ccb

V V Small ccb

− − Small rod

Blood, bone marrow

Blood, bone marrow, lymph node

V

Blood, cerebrospinal fluid, other

Blood, Wound, blood, lymph respiratory node aspirates, respiratory

−

−d

−

+

−

−

+

−

−

−

−

−

−

− 10:0, 14:0, 16:0, 3OH16:0, 18:1ω9c, 3-OH18:0

− 16:0, 18:1ω7c, 18:0, 19:0cycf

− 16:0, 17:0, 18:1ω7c

− 2-OH-12:0, 3OH-12:0, 16:1ω7c, 16:0, 18:1ω9c

− 14:0, 3-OH-14:0, 16:1ω7c, 16:0

− 16:0, 17:0, 3-OH14:0, 16:1ω7c, 18:1ω7c

− 16:1, 16:0; 14:0, 18:2, 3-OH-14:0

+ − Small ccb

a

+, greater than or equal to 90% positive; −, less than or equal to 10% positive; V, variable (11 to 89% positive); ccb, coccobacilli. Does not include Bartonella bacilliformis, which is the only motile species. Haemophilus spp. requiring X and V factors or V factor only. d While not required, X factor (hemin) enhances growth for many strains. e The number before the colon indicates the number of carbons; the number after the colon is the number of double bonds, “ω” indicates the location of the double bond counting from the hydrocarbon end of the carbon chain, “OH” indicates a hydroxy group at the 2 or 3 position from the carboxyl end, “c” indicates the cis isomer, and “cyc” indicates a cyclopropane ring structure. Hydroxy acids listed are at least 2% of the total cellular fatty acid (CFA) composition; all others are at least 10%. f B. canis lacks 19:0cyc. b c

TABLE 2

Characteristics of Francisella spp.a

Characteristic Gram stain (culture), safranin counterstain

Cell size (μm) Growth on blood agar

Cysteine heart blood agar, 48 h

Growth in NB (6% NaCl) Optimal growth temperature Catalase Oxidase Acid from: Glucoseb Maltose Sucrose Glycerol Citrulline ureidase Relative virulence (mice)

F. tularensis subsp. tularensis Faintly staining, pleomorphic, single, rarely chained, Gramnegative tiny coccobacilli 0.2–0.7 × 0.2 −

F. tularensis subsp. mediasiatica

F. tularensis subsp. holarctica

F. philomiragia

F. noatunensis

F. hispanensis

F. halioticida

F. guangzhouensis

As for F. tularensis subsp. tularensis

As for F. tularensis subsp. tularensis

As for F. tularensis subsp. tularensis

As for F. tularensis subsp. tularensis

Faintly staining, Gramnegative coccobacilli

Faintly staining, Gramnegative pleomorphic coccobacilli

Gram-negative coccobacilli

Gram-negative coccobacilli

0.2–0.7 × 0.2 −

0.2–0.7 × 0.2 −

0.7 × 1.7 +

0.7 × 1.7 +

NT −

0.5 × 1.5 +

0.5–1.0 NT; requires 70% artificial seawater for growth on Eugon agar Colonies observed 10– 14 days at 20°C were grayish to white, circular, slightly convex with entire margins

NT Delayed; weak

1- to 2-mm diam, raised, smooth colonies with entire margins; colonies display green tint and opalescent sheen −

As for F. tularensis subsp. tularensis

NT

As for F. tularensis subsp. tularensis but 2–4 mm in diam

Colonies are >4 mm in diam, creamy whitegray, mucoid, and smooth, with a purpletinted opalescent sheen

Colonies are low convex, white, slightly translucent and mucoid and ∼1 mm in diam after 4 days incubation at 22°C.

Colonies are convex, pale white to gray, 3- to 4-mm diameter

−

−

+

+

−

NT

+

NT

35–37°C

35–37°C

35–37°C

35–37°C

25 or 37°C

37°C

Weakly + −

Weakly + −

Weakly + −

Weakly + +

Weakly + +

20°C; no growth at 37°C + −

25-28 °C

Weakly + −

22°C; no growth at 37°C Weakly + −

+ −

+ + − + + High

+ + − − − Intermediate

− − + + + Intermediate

+ V + + + Low

Weakly + + V − NT None

V − − NT NT None

+ − + + NT NT

− NT − NT NT NT

+ + + − NT None

853

a

V, variable or slow reaction; NT, not tested; NB, nutrient broth. Delayed or variable reaction. F. tularensis subsp. mediasiatica does not ferment glucose (1).

b

F. novicida

Greenish gray mucoidopalescent colonies (∼2 mm)

854 n TABLE 3

BACTERIOLOGY Epidemiology of Francisella spp. affecting humansa

Francisella sp.

Infection source(s)

Geographic distribution (documented cases)

F. tularensis subsp. tularensis (type A)

Animals (primarily rabbits and cats), ticks, deerflies

North America (United States and Canada)

F. tularensis subsp. holarctica (type B) F. novicida

Animals (primarily rodents), ticks (mosquitoes in Europe) Salt water

Northern Hemisphere

F. philomiragia

Salt water

United States, Turkey, Switzerland, Australia

F. hispanensis Unclassified Francisella sp.

Unknown Unknown

Spain United States

United States, Thailand, Australia

Disease (humans) Tularemia (all forms); severe disease; A1b strains most virulent Tularemia (all forms); moderate to severe disease Very rare; mild illness (primarily compromised patients) Very rare; mild illness (primarily compromised patients) Very rare Very rare

a

Human infections documented in the published literature.

sphere. F. tularensis subsp. mediasiatica has been found only in regions of central Asia. More recently, the geographic distributions of F. tularensis subsp. holarctica and F. tularensis subsp. tularensis subpopulations have also been shown to differ. Subpopulations of F. tularensis subsp. holarctica within North America are distinct from subpopulations throughout the rest of the Northern Hemisphere, except Scandinavia (26). The F. tularensis subsp. tularensis A1a and A1b subpopulations predominate in the eastern half of the United States, whereas the A2 subpopulation appears to be restricted to the western United States (26, 27). Despite the broad geographic distribution of F. tularensis, overall disease occurrence is suspected to be low (28). Foci of endemicity have long been documented in regions of Russia and Kazakhstan, as well as Finland, Sweden, and the United States, where incidence may be higher (2). Annual cases are usually reported from most countries in Eastern Europe. Several outbreaks of tularemia have occurred over the last two decades in Spain, Sweden, Turkey, and Norway (29–33). In the United States, the incidence of tularemia has declined to 100 to 200 cases each year since the 1940s, when several thousand cases were reported annually (34). All states except Hawaii have reported cases. A total of 1,208 human cases from 44 states were reported to the Centers for Disease Control and Prevention (CDC) between 2000 and 2010. Four states, Arkansas, Missouri, Oklahoma, and Massachusetts, accounted for 48% of these cases (http://www.cdc.gov/ tularemia/statistics). Tularemia in humans is predominantly a rural disease; urban cases are rare. Most cases of tularemia occur in the summer months, with patients acquiring infection from tick or deerfly bites or from contact with infected animals (e.g., by skinning rabbits) (27, 34). Inhalation of an infective aerosol during landscaping and contact with infected cats are also important risk factors (27, 35). Tularemia cases are largely sporadic; outbreaks of tularemia occur rarely in the United States and may often follow tularemia epizootics in animals (30). An outbreak of pneumonic tularemia associated with landscaping activities occurred on Martha’s Vineyard in 2000 (35), and since that time, Martha’s Vineyard has recorded yearly cases (36). A more recent outbreak in the United States occurred in Utah in 2007 and was attributed to deerfly bites (37). From 1964 to 2004, 316 isolates recovered from human cases in the United States (39 different states) were submitted to the CDC; of these, 66% were F. tularensis subsp. tularensis and 34% were F. tularensis subsp. holarctica (38).

F. tularensis is a zoonotic pathogen, capable of infecting a large variety of animal species. The bacterium has been isolated from vertebrates, including mammals, amphibians, and birds, as well as from invertebrate arthropods (39–41). However, F. tularensis is most commonly associated with rodents and other mammals, particularly hares and rabbits. In both vertebrates and invertebrates, infection with F. tularensis can be lethal. In contrast to F. tularensis, F. novicida and F. philomiragia do not appear to circulate in animal or arthropod hosts. Only a single isolation of F. philomiragia from an animal host (muskrat) has been documented (6). Arthropods infected with F. novicida or F. philomiragia have never been identified. Transmission of F. novicida and F. philomiragia to humans appears to be associated with proximity to or contact with near-drowning in saltwater sources (8, 42). Most isolates of F. novicida are from North America, with a report of F. novicida-like organisms in Thailand and Australia (43, 44). Similarly, most isolates of F. philomiragia have been from North America, with two single incidences reported from Central and Eastern Europe and one from the Southern Hemisphere (Australia) (5, 8, 44–46). F. noatunensis been isolated from both the Northern and Southern Hemispheres (14, 15), whereas F. hispanensis, F. halioticida, and F. guangzhouensis have been recovered only in Spain, Japan, and China, respectively (11, 13, 16, 17).

CLINICAL SIGNIFICANCE F. tularensis Globally, tularemia is caused by two F. tularensis subspecies, F. tularensis subsp. tularensis (type A) and F. tularensis subsp. holarctica (type B). The disease has been known historically by a number of synonyms, such as rabbit fever, deerfly fever, market men’s disease, glandular type of tick fever, Ohara’s or yato-byo disease, and water rat trappers’ disease, attesting to the variety of clinical presentations, the infectious agent’s ubiquitous presence in nature, and the means by which humans may acquire the infection. The clinical spectrum of tularemia depends on the mode of transmission, the virulence of the infecting strain, the immune status of the host, and timely diagnosis and treatment (47). Tularemia can be misdiagnosed since the disease is rare and its symptoms are not unique; a sudden onset of

46. Francisella

chills, fever, headache, and generalized malaise characterize the onset of illness. The differential diagnosis includes a wide range of infectious diseases, such as cat scratch fever, mycobacterial infections, anthrax, brucellosis, legionellosis, and plague (47). Patients may present with any one of the clinical forms of tularemia: ulceroglandular, glandular, oculoglandular, oropharyngeal, typhoidal, and pneumonic (47). Upon infection, the period for disease onset is typically 3 to 5 days but can range from 1 to 21 days (28, 47). Disease presentation (i.e., signs and symptoms) correlates with the route of bacterial entry, with all forms of tularemia accompanied by fever (47). The most common form is ulceroglandular disease (45 to 80% of the reported cases), where the portal of entry is via an infective arthropod bite or other inoculation through the skin barrier. Glandular tularemia is similar to ulceroglandular disease but lacks the ulcerated site of infection. Oculoglandular tularemia occurs when the conjunctiva is the initial site of infection, usually as a result of the mechanical transfer of organisms from an infectious source to the eye by the fingers. Oropharyngeal tularemia occurs from ingestion of contaminated water or food and is associated with pharyngeal lymphadenopathy. Pneumonic tularemia occurs by direct inhalation of the organism and is considered the most severe form of the disease. Typhoidal tularemia is the most difficult form to recognize because there is no identified portal of entry and localized signs are absent. If untreated, bacterial dissemination from the primary sites of infection can lead to secondary clinical presentations, such as sepsis. The severity of infection can range from mild and selflimiting to fatal and is largely dependent on the infecting strain. Little to no tularemia-related mortality is reported in Europe and Asia, where only F. tularensis subsp. holarctica causes tularemia. In comparison, mortality in the United States ranges between 2.3% (those diagnosed by culture and serology) and 9% (culture-confirmed cases only) (38, 48), with tularemia caused by the more virulent F. tularensis subsp. tularensis as well as F. tularensis subsp. holarctica. In the last decade, molecular epidemiologic studies have demonstrated that among culture-confirmed human cases in the United States, infections due to A1b strains of F. tularensis subsp. tularensis result in a significantly higher rate of mortality (24%) than infections caused by F. tularensis subsp. tularensis A1a (4%) and A2 (0%) strains or F. tularensis subsp. holarctica (7%) (27). Virulence comparisons in mice corroborate these findings and show that A1b strains are more virulent than other F. tularensis subsp. tularensis strains (49). The third subspecies of F. tularensis, F. tularensis subsp. mediasiatica, has only been isolated from regions of Central Asia, and a description of human illness due to this subspecies is lacking in the literature. Experimental studies with rabbits indicate virulence comparable to that of F. tularensis subsp. holarctica (25).

F. novicida and F. philomiragia Human infections caused by F. novicida and F. philomiragia are very rare, with these organisms primarily infecting patients with underlying compromising conditions. Fewer than 20 cases of F. philomiragia infection have been described since the discovery of this species in 1974 (5, 8, 44–46). All but one case have involved a host with an impaired physical barrier to infection (near drowning) or an impaired immunologic defense system (chronic granulomatous disease or myeloproliferative disease). The drowning and water

n

855

exposure cases were associated with salt water and brackish water, in contrast to F. tularensis infections, which are associated with freshwater sources. Fewer than 10 cases of F. novicida infections have been reported worldwide (5, 8, 42– 44, 50, 51). Healthy individuals presented with mild illness (regional lymphadenopathy without fever), whereas patients with compromising conditions, primarily underlying liver disease, presented with fevers; in the latter cases, F. novicida was isolated from patient blood samples.

F. hispanensis Only one human case due to infection with F. hispanensis has been reported in the literature (11, 13). A previously healthy, immunocompetent patient presented with fever, myalgia, diarrhea, and lower left back pain; the organism was isolated from both blood and urine of this patient.

F. noatunensis and F. halioticida F. noatunensis and F. halioticida have never been associated with human infection, and the risk of human infection is considered very low. F. noatunensis is the significant cause of disease in farmed and wild fish, whereas F. halioticida has been associated with mortality in shellfish (giant farmed abalone) (52).

Another Francisella Species An unclassified Francisella sp. was isolated from the blood and cerebrospinal fluid of two different patients in the United States in 2005 and 2006 (19). No other cases with this species have been reported.

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS In the United States, F. tularensis is classified as a Tier 1 select agent. To receive or possess F. tularensis strains, laboratories must be registered with the Federal Select Agent Program (http://www.selectagents.gov). Clinical laboratories are exempt from the registration requirement with the Select Agent Program as pertains to conducting diagnostic testing for select agents. Both Tier 1 and non-Tier testing for select agents may be performed by clinical laboratories as long as the laboratory destroys or transfers any confirmed select agent (including clinical samples the isolate was derived from) with 7 days of identification. Laboratories identifying an organism as F. tularensis are required to report this finding immediately; laboratories need to complete Form 4 and submit it to the Select Agent Program. If the organism is to be transferred following identification, then the laboratory must also complete Form 2 and obtain transfer approval from the Select Agent Program. Select agent report forms, contact information, laboratory registration information, and pertinent citations of the U.S. Federal Code may be found at http:// www.selectagents.gov. Personnel handling diagnostic cultures of F. tularensis are at considerable risk for infection. Due to the extremely low infectious dose for F. tularensis, tularemia has been one of the most commonly reported laboratory-associated bacterial infections (53, 54). Even though the use of biological safety cabinets and prophylactic antibiotic therapy (as well as vaccination, where available) provides safeguards for laboratory workers, these precautions have not fully eliminated laboratory exposures or modified practices in the clinical laboratory to minimize risks (55). All patient specimens should be handled and processed in biosafety level 3 (BSL3) or BSL2 with BSL3 precautions,

856 n

BACTERIOLOGY

wearing gloves and lab coats or gowns and working in a class II biosafety cabinet, and clinical laboratories should be aware of the sentinel-level clinical microbiology laboratory guidelines as outlined by the American Society for Microbiology (http://www.asm.org/index.php/issues/sentinel-lab oratory-guidelines). Aerosol inhalation during manipulation of cultures presents the greatest risk to laboratory workers, due to the high concentration of organisms. BSL3 practices, containment equipment, and facilities are recommended for all manipulations of suspicious cultures (56). Preparatory work on cultures for automated identification systems should be performed at BSL3. Laboratories may want to consider developing policies that encourage physicians to alert the laboratory if a diagnosis of tularemia is expected. The choice of specimen for diagnostic testing is generally dependent on the form of clinical illness: ulceroglandular, glandular, oculoglandular, oropharyngeal, pneumonic, or typhoidal tularemia (47). Whole blood is an acceptable specimen for all clinical forms of tularemia, although this sample may be negative, particularly if the disease is in the early stages of progression. Serum for antibody detection is a standard specimen taken for diagnosis of all forms of illness. The first specimen should be collected as early in the course of infection as possible, followed by a second specimen taken in the convalescent period (at least 14 days apart and preferably 3 to 4 weeks after onset of symptoms). Pharyngeal swabs, bronchial or tracheal washes or aspirates, sputum, transthoracic lung aspirates, and pleural fluid are appropriate specimens for pneumonic, typhoidal, or oropharyngeal tularemia. Swabs of visible lesions or affected areas have been used most commonly for ulceroglandular and oculoglandular tularemia. Aspirates from lymph nodes or lesions can be used for diagnosis of ulceroglandular, glandular, and oropharyngeal tularemia. Necropsied materials from animals that are appropriate for testing include samples from visible abscesses as well as samples from lymph node, lung, liver, and spleen tissues and bone marrow. For specimens to be tested by culture, it is important where possible to decontaminate the surface area prior to specimen collection, since contamination of the sample with normal flora could interfere with the interpretation of culture results. To minimize loss in viability, specimens should be delivered to the laboratory within 24 h and preferably within 2 h. In general, if transport is >24 h, specimens should be stored chilled (2 to 8°C) in an appropriate medium until processed in the laboratory. Freezing of samples, unless in a preservative environment, such as tissue specimens or glycerol-containing solutions, is not recommended because of the lysis of live bacteria upon thawing. Swabs should be placed in Amies agar with charcoal, a commercial transport system designed for anaerobic and aerobic pathogens (Becton, Dickinson and Company, Franklin Lakes, NJ). F. tularensis should remain viable for 7 days at ambient room temperature when stored in Amies medium (57). Stuart medium, designed for transporting gonococcal specimens, and saline are inadequate for keeping F. tularensis viable during transport (57). For serum samples, separation from blood should take place as soon as possible, preferably within 24 h. Sera may be stored at 2 to 8°C for up to 10 days. If testing is delayed for a long period, serum samples may be frozen. Guanidine isothiocyanate-containing buffer, which preserves F. tularensis DNA for up to 1 month, is recommended if there is a delay in PCR testing of samples (57). Arthropods may be stored intact in 2% NaCl for culture analysis or in ethanol for molecular testing.

DIRECT EXAMINATION Microscopy Fresh clinical specimens (ulcer and wound swabs, tissues, and aspirates) where the concentration of organisms might be expected to be high can be directly examined by microscopy by Gram straining, direct fluorescent-antibody (DFA) binding, or immunohistochemistry (IHC). Under microscopic examination of Gram-stained specimens, Francisella cells (single and pleomorphic) appear tiny and counterstain so faintly with safranin that they can easily be missed (Fig. 1). Basic fuchsin counterstains F. tularensis better than safranin. Due to the small size of F. tularensis, Gram staining of clinical specimens is usually of little diagnostic value. F. tularensis should be included in the clinical laboratory differential if very small, faintly staining Gram-negative coccobacilli are seen. F. tularensis cells are smaller than Haemophilus.

Antigen Detection A rapid and specific staining method (DFA) for detection of F. tularensis in specimens includes the use of a fluorescein isothiocyanate-labeled hyperimmune rabbit polyclonal antibody directed against whole, killed F. tularensis cells. This staining method can be used to presumptively identify F. tularensis subsp. tularensis and F. tularensis subsp. holarctica in clinical specimens (47). Cultures, lesions, tissues, or aspirates may be rapidly assessed by this approach. This DFA reagent does not react well or at all with F. novicida or F. philomiragia. IHC staining using a monoclonal antibody directed against the lipopolysaccharide has been used successfully to visualize F. tularensis in formalin-fixed tissues (58). Neither the DFA reagent nor the IHC reagent is commercially available. DFA testing of specimens is provided by reference laboratories (Laboratory Response Network [LRN]) in the United States.

Nucleic Acid Detection Because of the relative rarity of human tularemia, evaluation of molecular diagnostics with clinical specimens has been challenging. PCR-based diagnostic methods have been used most commonly for diagnosis of ulceroglandular tularemia, the most prevalent clinical form. DNA detection by conventional PCR directed at the tul4 gene (unique to Francisella

FIGURE 1 Gram stain of F. tularensis. Magnification, ∼×810. doi:10.1128/9781555817381.ch46.f1

46. Francisella n 857

spp.) has been successfully and widely applied for diagnosis of ulceroglandular tularemia (47, 57, 59). The tul4 PCR assay displays a sensitivity of 75% when applied to wound specimens from patients with ulceroglandular tularemia and was shown to be more sensitive than culture (sensitivity of 62%) (57). Real-time PCR assays using the TaqMan 5′ nuclease assay have been applied to detect F. tularensis DNA in a variety of clinical specimens, including ulcer specimens, aspirates, pharyngeal swabs, lymph node specimens, bronchial washes, and pleural fluid (60–63). These assays target multiple Francisella genes (ISFtu2 element, iglC, tul4, and fopA genes). Real-time PCR assays for diagnosis of tularemia are available in LRN reference laboratories in the United States. A limitation of most PCR-based diagnostics for F. tularensis is the inability to discriminate F. tularensis from F. novicida, due to their high degree of genetic relatedness. While this may not be significant for patient management, the correct identification of species has biothreat, epidemiologic, and public health importance. A recent study evaluated 38 published PCR assays against the current known diversity of the Francisella genus, and it found that most nucleic acid assays are not specific for only a single Francisella species (64). Detection of F. tularensis in ectoparasites is based primarily on PCR methods. Of note, FLEs, present in a wide range of tick species, have been shown to cross-react with molecular targets used for the detection of F. tularensis, leading to falsepositive results (20, 65, 66). If molecular assays are to be used for screening environmental samples (water or soil) for F. tularensis, it is important to evaluate the assays for cross-reactivity with other Francisellaceae members, particularly F. novicida, present in these sample types (18, 67). 16S rRNA gene sequencing can be used to discriminate F. tularensis from FLEs or environmental Francisella spp. other than F. novicida (65, 67).

Isolation Procedures F. tularensis is slow growing and fastidious; it is auxotrophic for cysteine and consequently requires an exogenous source of cysteine in order to grow well on artificial medium. F. tularensis grows on several media common to clinical laboratories, including chocolate agar (CA) (Fig. 2A), buffered charcoalyeast extract agar (BCYE), and Thayer-Martin agar. It also grows in thioglycolate broth, although growth in broth is not recommended in clinical laboratories due to the increased risk of exposure via aerosolization. When supplemented with 1 to 2% IsoVitaleX (which contains cysteine and other supplements), general bacteriological media (tryptic soy broth

and Mueller-Hinton broth) can also support the growth of F. tularensis. The organism grows slowly (60-min generation time); good growth, therefore, is obtained by prolonged incubation (48 h or longer). Suspicious cultures should be incubated at 35 to 37°C aerobically and observed daily for up to 14 days; CO2 neither impedes nor is required for growth. A specialized medium, cysteine heart agar supplemented with 9% chocolatized sheep blood (CHAB), is often used for the growth of F. tularensis in reference laboratories (47). F. tularensis grows well on CHAB, as this is a high-nutrient medium. Additionally, F. tularensis displays distinctive colony morphology on CHAB which can aid in identification (Fig. 2B). F. tularensis grows poorly or not at all on general bacteriological media, such as sheep blood agar. Nutritionally enriched specimens, such as blood or tissue, provide an intrinsic source of cysteine that may initially permit F. tularensis growth on general bacteriological media. Upon subculture, the fastidious nature of F. tularensis will become evident as the exogenous compounds are depleted, leading to the loss of the bacterium’s viability unless the subculture is propagated on cysteine-supplemented medium. Clinical samples from normally sterile sites can be plated on nonselective agars that support the growth of F. tularensis. Care should be taken not to permit laboratory contamination of F. tularensis isolates by environmental bacteria such as Staphylococcus, as these bacteria rapidly outcompete and inhibit the growth of F. tularensis on nonselective media (68). Clinical specimens obtained from nonsterile sites or autopsy and environmental sources should be plated on antibiotic-containing media. Incorporation of an antibiotic supplement (7.5 mg/liter of colistin, 2.5 mg/liter of amphotericin, 0.5 mg/liter of lincomycin, 4.0 mg/liter of trimethoprim, and 10 mg/liter of ampicillin) into the medium has been demonstrated to prevent other organisms from overwhelming F. tularensis (68). Commercially available antibiotic-containing media that support the growth of F. tularensis include improved Thayer-Martin agar (Remel, Lenexa, KS), BCYE selective agar containing polymyxin B, anisomycin, and vancomycin (Remel or Becton, Dickinson), and cysteine heart agar containing antibiotics (polymyxin B and penicillin) (Remel). Specimens for culture should be taken on the basis of clinical presentation and before administration of antibiotics. Fresh clinical material that is likely to contain high concentrations of F. tularensis organisms, such as ulcer and wound specimens and lymphoid tissue (liver, spleen, or affected lymph node tissue), is inoculated directly onto an agar plate by using a sample-laden swab or bacteriological loop. Larger inocula are necessary for the recovery of F. tularensis from specimens that contain a lower concentration of the organisms, such as aspirates of pharyngeal washes, bronchial wash fluids, pleural fluids, and environmental samples. Whole blood should be directly inoculated into blood culture bottles (47), with subsequent culture of blood culture-positive specimens on agar. Blood culture bottles should be incubated for 5 to 7 days to allow for growth of F. tularensis.

IDENTIFICATION

FIGURE 2 F. tularensis on CA (A) and CHAB (B) after 72 and 48 h of growth, respectively. (Panel A is courtesy of the CDC Public Health Image Library [Larry Stauffer, Oregon State Public Health Laboratory].) doi:10.1128/9781555817381.ch46.f2

Because of the rarity as well as the sporadic nature of most tularemia cases, the organism is often not easily identified when it is cultured. The isolation of a very tiny (individual cells may be difficult to discern), poorly counterstaining (by safranin), slow-growing Gram-negative coccobacillus (Fig. 1) that produces 1- to 2-mm gray to gray-white colonies on CA after 48 h, scant to no growth

858 n BACTERIOLOGY

on blood agar, and no growth on MacConkey agar should raise suspicion for F. tularensis. F. tularensis is oxidase negative, weakly catalase positive, β-lactamase positive, X and V or satellite growth negative, and urease negative. Gram stain, oxidase, XV or satellite, and urease tests can help differentiate F. tularensis from other similar Gramnegative organisms, including Brucella spp., Haemophilus influenzae, Bartonella spp., Yersinia pestis, Pasteurella multocida, and Acinetobacter spp. (Table 1). If F. tularensis is suspected based on these results, the isolate should be sent to a reference laboratory that can confirm (or rule out) its identification as F. tularensis. In the United States, all states have at least one reference laboratory that is part of the LRN. LRN laboratories are able to confirm the identification of bacterial select agents, including F. tularensis. See also http://www.bt.cdc.gov/lrn. The colony morphology of F. tularensis is most distinctive when the organism is grown on CHAB; after 48 h at 37°C, F. tularensis colonies are 1 to 2 mm, raised, shiny, green, and opalescent (Fig. 2B). On CA, BCYE, and Thayer-Martin agar after 48 h at 37°C, F. tularensis colonies are ∼1 mm, white to gray, and smooth and opaque and have an entire margin (Fig. 2A). F. novicida and F. philomiragia grow more robustly than F. tularensis (Table 2), and colonies of F. philomiragia are mucoid. Isolates can be identified as F. tularensis using antigen or molecular detection methods, including slide agglutination, DFA staining (Fig. 3), PCR, or sequencing. The slide agglutination test identifies a suspicious culture by mixing commercially available (Becton, Dickinson and Company) polyclonal rabbit anti-F. tularensis antibody with suspicious cultures. DFA staining can also be used to identify F. tularensis subsp. tularensis and F. tularensis subsp. holarctica isolates (47). Care should be taken to ensure that prepared smears are not too thick, as this can interfere with the performance and interpretation of the test. In general, antigen-based identification methods work optimally when fresh cultures (24 h) are tested. If a culture older than 24 h is to be tested by antigen detection methods, a fresh subculture should be prepared. Polyclonal antibodies to whole killed F. tularensis, used in both the DFA and slide agglutination tests, generally react poorly or not at all with F. novicida due to differences in the O antigens of the lipopolysaccharides of the two organisms (47, 69). Thus, this method for identification of F. tularensis isolates is preferred, due to this specificity. PCR methods targeting F. tularensis-specific genes can also be used for the identification of suspicious cultures (59, 61). As described earlier in this chapter, it is important to

consider that many F. tularensis PCR assays cross-react with F. novicida. Therefore, it may be important to rule out F. novicida as the cause of infection, particularly in areas where tularemia has not been previously reported. 16S rRNA gene sequencing, using universal 16S rRNA gene primers or Francisella-specific 16S rRNA gene primers, can identify the organism as a Francisella species (10, 29, 70). Due to the high degree of genetic relatedness between F. tularensis and F. novicida, 16S rRNA PCR and sequencing do not readily distinguish these two species. F. novicida isolates have been misidentified as F. tularensis in clinical laboratories by 16S sequencing (42). Members of the Francisella genus have a unique fatty acid composition, high in saturated even-chain acids (C10:0, C14:0, and C16:0) and two long-chain hydroxyl acids (3-OH-C16:0 and 3-OH-C18:0) (71), which can be used to identify the organism as belonging to the Francisella genus (Table 1) (1, 5, 21). Bacterial identification systems that rely on fatty acid profiles have been used to identify F. tularensis; however, it is important to note that commercial systems do not include F. novicida or many other Francisella spp. in their databases. Indeed, misidentification of F. novicida as F. tularensis, in a clinical laboratory setting, using these systems has been described (42). Commercial biochemical identification systems available in clinical laboratories are not recommended for the diagnosis of F. tularensis because of the high probability of misidentification and the potential for generating infectious aerosols (47). Misidentification is largely due to a lack of biochemical reactivity by F. tularensis and a limited number of strains used in the development of databases. Because of the limitations in identifying F. tularensis isolates using automated systems in clinical laboratories, a high level of suspicion by clinical microbiologists is essential for accurate and timely diagnosis of tularemia. Results from automated identification systems should be analyzed critically in the clinical laboratory, taking into consideration the reliability of these systems for accurate identification of slow-growing and/or rare organisms. Potential indications of misidentification by automated systems include a low probability or confidence value and/or an identification of a pathogen inconsistent with the clinical or microbiology picture. The utility of matrix-assisted laser desorption ionization– time of flight (MALDI-TOF) mass spectrometry (MS) combined with dedicated bioinformatics and statistical methods for identification and differentiation of Francisella strains was recently demonstrated (72). Forty-four strains of F. tularensis were accurately identified to the subspecies level as well as differentiated from six strains of F. philomiragia (72). Thus, this method shows promise as a method for future identification of F. tularensis in the clinical laboratory. At the current time, however, documented use of MALDITOF MS with commercially available databases in clinical laboratory settings is lacking.

TYPING SYSTEMS

FIGURE 3 DFA staining of a culture of F. tularensis. Magnification, ∼×490. doi:10.1128/9781555817381.ch46.f3

Once an isolate has been identified as a Francisella sp., supplemental tests can be used for additional characterization, including typing of species, subspecies, and strain. Oxidase can be used to differentiate F. philomiragia from F. novicida and F. tularensis (Table 2). Glycerol fermentation and citrulline ureidase activity distinguish F. tularensis subsp. tularensis from F. tularensis subsp. holarctica (Table 2); conventional assays for these biochemical tests have been described (73). The 96-well automated MicroLog MicroStation system with GN2 microplates (Biolog Inc., Hayward, CA) can also be used to

46. Francisella

assess the glycerol fermentation of F. tularensis (47). The genus can be typed by sequence analysis of the 16S rRNA gene (10, 70). PCR methods (both conventional and realtime) can type isolates at the level of genus, species, subspecies (F. tularensis subsp. tularensis or F. tularensis subsp. holarctica), and subclade (F. tularensis subsp. tularensis subclades A1 and A2) (47, 74). Pulsed-field gel electrophoresis can differentiate the three F. tularensis subsp. tularensis subpopulations, A1a, A1b, and A2 (27). For discrimination of individual strains, an optimized, multiplex multilocus variable-number tandem-repeat assay for F. tularensis, based on 11 different repeats in the genome, can be used (75, 76).

SEROLOGIC TESTS A confirmatory diagnosis of tularemia can be established serologically by demonstrating a 4-fold or significant change in specific antibody titers to or response between acute- and convalescent-phase sera to F. tularensis antigen (47). A single positive antibody titer for a patient with no history of tularemia vaccination is a presumptive diagnosis of tularemia. As antibody responses to F. tularensis generally require 10 to 20 days to develop postinfection, serologic testing is of limited use in clinical management of patients with acute illness. IgM, IgA, and IgG antibodies may appear simultaneously (77, 78), and antibodies to F. tularensis can persist for more than 10 years (79, 80). IgM responses have been shown to be sustained for long periods and are therefore not indicative of early or recent infection (77, 79). Agglutination testing, either by the tube agglutination (TA) or the microagglutination (MA) method, is a standard serologic test for determining the presence of antibodies in patients with tularemia (47, 79, 81). Formalin-killed antigen (prepared from F. tularensis subsp. holarctica strain LVS) is commercially available from Becton, Dickinson, and is also prepared within reference laboratories worldwide. In the United States, a single specimen with a TA titer of ≥1:160 or an MA titer of ≥1:128 is considered positive. Formalin-killed F. tularensis whole-cell antigen may display low-level crossreactivity with Brucella antibodies (82, 83). No crossreactivity of F. novicida or F. philomiragia sera has been observed with F. tularensis-killed cells (9). Enzyme-linked immunosorbent assays (ELISAs) have been adopted for use in the parts of Europe where tularemia is endemic (47, 79, 84, 85). The lipopolysaccharide and/or outer membrane fraction remains the primary ELISA antigen used in test applications. Antigenic differences between F. tularensis subsp. tularensis and F. tularensis subsp. holarctica have not been identified for use in serology assays. Thus, serology assays do not distinguish the infecting subspecies. This is of the most importance in North America, where both F. tularensis subsp. tularensis and F. tularensis subsp. holarctica cause tularemia. F. tularensis organisms are intracellular bacteria and are capable of eliciting both humoral and cell-mediated immunity (77). The latter response has been known to remain strong 25 years after infection (86). Host T cells retain proliferative responses to unique F. tularensis membrane proteins, with concomitant increases in interferon and interleukin-2 levels (77, 86–88). Tests for measuring the cellmediated immune response are specialized and are not routinely used for diagnosis of tularemia (89).

ANTIMICROBIAL SUSCEPTIBILITIES F. tularensis infections are treatable with narrow-spectrum antibiotics. Naturally occurring resistance in F. tularensis to aminoglycosides, tetracyclines, and fluoroquinolones has

n

859

never been shown (47, 90–92). The risk for development of antibiotic resistance in clinical disease is low, as tularemia is an end-stage disease and also is not transmitted from person to person (47). Treatment failures in tularemia patients correlate with a delay in the initiation of antibiotics with respect to symptom onset, as opposed to development of resistance (60, 93, 94). Lymph node suppuration, which is nonresponsive to all classes of antibiotics and requires surgical drainage, can develop in these cases. Considerably longer recovery times are observed in these cases and can be greater than 70 days (94). β-Lactam antibiotics are not used for treatment of tularemia, as F. tularensis strains encode a class A β-lactamase, FTU-1 (95). β-Lactamase resistance appears to be limited to penicillins; in vitro MIC determinations indicate that FTU1 does not confer resistance to other β-lactams, including first- and second-generation cephalosporins (95). Although third-generation cephalosporins have been shown to be active against F. tularensis in vitro, clinical experience indicates that ceftriaxone is not effective for treatment of tularemia (96). Macrolides are also not recommended for treatment of tularemia, as type B strains from Central and Eastern Europe and Asia are naturally resistant to erythromycin and other macrolides (97). Antimicrobial susceptibility testing of F. tularensis is not performed in clinical microbiology laboratories because of safety concerns in working with this organism and because resistance to antibiotics used for clinical treatment of tularemia has never been reported (90). The Clinical and Laboratory Standards Institute (CLSI) has published interpretative criteria and quality control limits for broth microdilution of F. tularensis using Mueller-Hinton medium supplemented with 2% IsoVitaleX (98, 99).

EVALUATION, INTERPRETATION, AND REPORTING OF RESULTS Serology is a common method for laboratory confirmation of F. tularensis infection, due largely the organism’s slow growth and fastidious nature. Nonetheless, culture provides a conclusive diagnosis of infection and whenever possible should be attempted using appropriate biosafety measures. A very tiny Gram-negative bacterium that grows slowly, shows fastidious growth characteristics, and is oxidase negative, weakly catalase positive, urease negative, X/V or satellite negative, and β-lactamase positive should be strongly suspected as F. tularensis and referred to a reference laboratory. Confirmation of F. tularensis infections includes (i) identification of a culture as F. tularensis and/or (ii) a 4-fold difference in titers in acuteand convalescent-phase serum samples, with one of the paired samples having a positive titer. A positive test result for a primary clinical specimen using antigen or molecular detection methods, including DFA staining, IHC staining, or PCR, provides only a presumptive diagnosis of F. tularensis. A single positive serum sample is also considered presumptive for tularemia. For all cases presumed to be tularemia, it is necessary to verify that the patients’ symptoms are compatible with tularemia. In the United States, tularemia is a nationally notifiable disease and F. tularensis is also classified as a select agent. Identification of a bacterial select agent requires isolation of the organism in culture and confirmation. Laboratories identifying an organism as F. tularensis are required to report this finding immediately; laboratories need to complete Form 4 and submit it to the Select Agent Program. Report forms, contact information, laboratory registration information, and pertinent citations of the U.S. Federal Code may be found at http://www.selectagents.gov.

860 n

BACTERIOLOGY

REFERENCES 1. Sjöstedt A. 2005. Francisella, p 200–210. In Brenner DJ, Krieg NR, Staley JT, Garrity GM (ed), Bergey’s Manual of Systematic Bacteriology, 2nd ed, vol 2. The Proteobacteria. Springer-Verlag, New York, NY. 2. Sjöstedt A. 2007. Tularemia: history, epidemiology, pathogen physiology, and clinical manifestations. Ann N Y Acad Sci 1105:1–29.165. 3. Sjödin A, Svensson K, Ohrman C, Ahlinder J, Lindgren P, Duodu S, Johansson A, Colquhoun DJ, Larsson P, Forsman M. 2012. Genome characterisation of the genus Francisella reveals insight into similar evolutionary paths in pathogens of mammals and fish. BMC Genomics 13:268. 4. Larsson P, Elfsmark DK, Svensson K, Wikström P, Forsman M, Brettin T, Keim P, Johansson A. 2009. Molecular evolutionary consequences of niche restriction in Francisella tularensis, a facultative intracellular pathogen. PLoS Pathog 5:e1000472. 5. Hollis DG, Weaver RE, Steigerwalt AG, Wenger JD, Moss CW, Brenner DJ. 1989. Francisella philomiragia comb. nov. (formerly Yersinia philomiragia) and Francisella tularensis biogroup novicida (formerly Francisella novicida) associated with human disease. J Clin Microbiol 27:1601–1608. 6. Jensen WI, Owen CR, Jellison WL. 1969. Yersinia philomiragia sp. n., a new member of the Pasteurella group of bacteria, naturally pathogenic for the muskrat (Ondatra zibethica). J Bacteriol 100:1237–1241. 7. Larson CL, Wicht W, Jellison WL. 1955. A new organism resembling P. tularensis isolated from water. Public Health Rep 70:253–258. 8. Wenger JD, Hollis DG, Weaver RE, Baker CN, Brown GR, Brenner DJ, Broome CV. 1989. Infection caused by Francisella philomiragia (formerly Yersinia philomiragia): a newly recognized human pathogen. Ann Intern Med 110:888–892. 9. Owen CR, Buker EO, Jellison WL, Lackman DB, Bell JF. 1964. Comparative studies of Francisella tularensis and Francisella novicida. J Bacteriol 87:676–683. 10. Forsman M, Sandström G, Sjöstedt A. 1994. Analysis of 16S ribosomal DNA sequences of Francisella strains and utilization for determination of the phylogeny of the genus and for identification of strains by PCR. Int J Syst Bacteriol 44:38–46. 11. Huber B, Escudero R, Busse HJ, Seibold E, Scholz HC, Anda P, Kämpfer P, Splettstoesser WD. 2010. Description of Francisella hispaniensis sp. nov., isolated from human blood, reclassification of Francisella novicida (Larson et al. 1955) Olsufiev et al. 1959 as Francisella tularensis subsp. novicida comb. nov. and emended description of the genus Francisella. Int J Syst Evol Microbiol 60:1887–1896. 12. Johansson A, Celli J, Conlan W, Elkins KL, Forsman M, Keim PS, Larsson P, Manoil C, Nano FE, Petersen JM, Sjöstedt A. 2010. Objections to the transfer of Francisella novicida to the subspecies rank of Francisella tularensis. Int J Syst Evol Microbiol 60:1717–1718. 13. Escudero R, Elía M, Sáez-Nieto JA, Menéndez V, Toledo A, Royo G, Rodríguez-Vargas M, Whipp MJ, Gil H, Jado I, Anda P. 2010. A possible novel Francisella genomic species isolated from blood and urine of a patient with severe illness. Clin Microbiol Infect 16:1026–1030. 14. Mikalsen J, Olsen AB, Tengs T, Colquhoun DJ. 2007. Francisella philomiragia subsp. noatunensis subsp. nov., isolated from farmed Atlantic cod (Gadus morhua L.). Int J Syst Evol Microbiol 57:1960–1965. 15. Ottem KF, Nylund A, Karlsbakk E, Friis-Møller A, Kamaishi T. 2009. Elevation of Francisella philomiragia subsp. noatunensis Mikalsen et al. (2007) to Francisella noatunensis comb. nov. [syn. Francisella piscicida Ottem et al. (2008) syn. nov.] and characterization of Francisella noatunensis

16.

17.

18.

19.

20.

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

subsp. orientalis subsp. nov., two important fish pathogens. J Appl Microbiol 106:1231–1243. Brevik OJ, Ottem KF, Kamaishi T, Watanabe K, Nylund A. 2011. Francisella halioticida sp. nov., a pathogen of farmed giant abalone (Haliotis gigantea) in Japan. J Appl Microbiol 111:1044–1056. Qu PH, Chen SY, Scholz HC, Busse HJ, Gu Q, Kämpfer P, Foster JT, Glaeser SP, Chen C, Yang ZC. 2013. Francisella guangzhouensis sp. nov., isolated from air conditioning systems. Int J Syst Evol Microbiol 63:3628–3635. Petersen JM, Carlson J, Yockey B, Pillai S, Kuske C, Garbalena G, Pottumarthy S, Chalcraft L. 2009. Direct isolation of Francisella spp. from environmental samples. Lett Appl Microbiol 48:663–667. Kugeler KJ, Mead PS, McGowan KL, Burnham JM, Hogarty MD, Ruchelli E, Pollard K, Husband B, Conley C, Rivera R, Kelesidis T, Lee WM, Mabey W, Winchell JM, Stang HL, Staples JE, Chalcraft LJ, Petersen JM. 2008. Isolation and characterization of a novel Francisella sp. from human cerebrospinal fluid and blood. J Clin Microbiol 46:2428–2431. Scoles GA. 2004. Phylogenetic analysis of the Francisellalike endosymbionts of Dermacentor ticks. J Med Entomol 41:277–286. Weyant RS, Moss CW, Weaver RE, Hollis DG, Jordan JG, Cook EC, Daneshvar MI. 1996. Identification of Unusual Pathogenic Gram-Negative Aerobic and Facultatively Anaerobic Bacteria, 2nd ed. The Williams & Wilkins Co, Baltimore, MD. Meshcheriakova IS, Kormilitsyna MI, Rodionova IV, Konstantinova ND. 1995. The characteristics of new species of pathogenic microorganisms in the genus Francisella. Zh Mikrobiol Epidemiol Immunobiol 5:3–8. (In Russian.) Olsufiev NG, Emelyanova OS, Dunaeva TN. 1959. Comparative study of strains of B. tularense in the old and new world and their taxonomy. J Hyg Epidemiol Microbiol Immunol 3:138–149. Mikalsen J, Olsen AB, Rudra H, Moldal T, Lund H, Djønne B, Bergh O, Colquhoun DJ. 2009. Virulence and pathogenicity of Francisella philomiragia subsp. noatunensis for Atlantic cod, Gadus morhua L., and laboratory mice. J Fish Dis 32:377–381. Olsufjev NG, Meshcheryakova IS. 1982. Infraspecific taxonomy of tularemia agent Francisella tularensis McCoy et Chapin. J Hyg Epidemiol Microbiol Immunol 3:291–299. Molins CR, Petersen JM. 2010. Subpopulations of F. tularensis subspecies tularensis and holarctica: identification and associated epidemiology. Future Microbiol 5:649–661. Kugeler KJ, Mead PS, Janusz AM, Staples JE, Kubota KA, Chalcraft LG, Petersen JM. 2009. Molecular epidemiology of Francisella tularensis in the United States. Clin Infect Dis 48:863–870. Dennis DT, Inglesby TV, Henderson DA, Bartlett JG, Ascher MS, Eitzen E, Fine AD, Friedlander AM, Hauer J, Layton M, Lillibridge SR, McDade JE, Osterholm MT, O’Toole T, Parker G, Perl TM, Russell PK, Tonat K, Working Group on Civilian Biodefense. 2001. Tularemia as a biological weapon: medical and public health management. JAMA 285:2763–2773. Anda P, del Pozo JS, Garcia JMD, Escudero R, Peña FJG, Velasco MCL, Sellek RE, Chillarón MRJ, Serrano LPS, Navarro JFM. 2001. Waterborne outbreak of tularemia associated with crayfish fishing. Emerg Infect Dis 7:575– 582. Martín CM, Gallardo T, Mateos L, Vián E, García MJ, Ramos J, Berjón AC, del Carmen Viña M, García MP, Yáñez J, González LC, Muñoz T, Allue M, Andrés C, Ruiz C, Castrodeza J. 2007. Outbreak of tularaemia in Castilla y León, Spain. Euro Surveill 12:E071108.1. Akalin H, Helvaci S, Gedikogˇ lu S. 2009. Re-emergence of tularemia in Turkey. Int J Infect Dis 13:547–551.

46. Francisella 32. Larssen KW, Afset JE, Heier BT, Krogh T, Handeland K, Vikøren T, Bergh K. 2011. Outbreak of tularaemia in central Norway, January to March 2011. Euro Surveill 16(13):pii=19828. 33. Payne L, Arneborn M, Tegnell A, Giesecke J. 2005. Endemic tularemia, Sweden, 2003. Emerg Infect Dis 11:1440–1442. 34. Centers for Disease Control and Prevention. 2002. Tularemia—United States, 1990–2000. MMWR Morb Mortal Wkly Rep 51:182–184. 35. Feldman KA, Enscore R, Lathrop S, Matyas B, McGuill M, Schriefer M, Stiles-Enos D, Dennis D, Hayes E. 2001. Outbreak of primary pneumonic tularemia on Martha’s Vineyard. N Engl J Med 345:1601–1606. 36. Matyas BT, Nieder HS, Telford SR III. 2007. Pneumonic tularemia on Martha’s Vineyard: clinical, epidemiologic, and ecological characteristics. Ann N Y Acad Sci 1105:351– 377. 37. Petersen JM, Carlson JK, Dietrich G, Eisen RJ, Coombs J, Janusz AM, Summers J, Beard CB, Mead PS. 2008. Multiple Francisella tularensis subspecies and clades, tularemia outbreak, Utah. Emerg Infect Dis 14:1928–1930. 38. Staples JE, Kubota KA, Chalcraft LG, Mead PS, Petersen JM. 2006. Epidemiologic and molecular analysis of human tularemia, United States, 1964–2004. Emerg Infect Dis 12:1113–1118. 39. Bell JF. 1980. Tularemia, p 161–193. In Steele JH (ed), CRC Handbook Series in Zoonoses. CRC Press, Boca Raton, FL. 40. Hopla CE, Hopla AK. 1994. Tularemia, p 113–126. In Beran GW, Steele JH (ed), Handbook of Zoonoses, 2nd ed. CRC Press, Boca Raton, FL. 41. Jellison WL, Owen CR, Bell JF, Kohls GM. 1961. Tularemia and animal populations: ecology and epi-zootiology. Wildl Dis 17:1–15. 42. Brett M, Doppalapudi A, Respicio-Kingry LB, Myers D, Husband B, Pollard K, Mead P, Petersen JM, Whitener CJ. 2012. Francisella novicida bacteremia after a neardrowning accident. J Clin Microbiol 50:2826–2829. 43. Leelaporn A, Yongyod S, Limsrivanichakorn S, Yungyuen T, Kiratisin P. 2008. Francisella novicida bacteremia, Thailand. Emerg Infect Dis 14:1935–1937. 44. Whipp MJ, Davis JM, Lum G, de Boer J, Zhou Y, Bearden SW, Petersen JM, Chu MC, Hogg G. 2003. Characterization of a novicida-like subspecies of Francisella tularensis in Australia. J Med Microbiol 52:839–842. 45. Friis-Møller A, Lemming LE, Valerius NH, Bruun B. 2004. Problems in identification of Francisella philomiragia associated with fatal bacteremia in a patient with chronic granulomatous disease. J Clin Microbiol 42:1840–1842. 46. Sicherer SH, Asturias EJ, Winkelstein JA, Dick JD, Willoughby RE. 1997. Francisella philomiragia sepsis in chronic granulomatous disease. Pediatr Infect Dis J 16:420– 422. 47. World Health Organization. 2007. WHO Guidelines on Tularemia. World Health Organization, Geneva, Switzerland. http://www.cdc.gov/tularemia/resources/whotulare miamanual.pdf. 48. Evans ME, Gregory DW, Schaffner W, McGee ZA. 1985. Tularemia: a 30-year experience with 88 cases. Medicine (Baltimore) 64:251–269. 49. Molins CR, Delorey M, Yockey BM, Young JW, Sheldon SW, Reese SM, Schriefer ME, Petersen JM. 2010. Virulence comparison among Francisella tularensis subsp. tularensis clades in mice. PLoS One 5(4):e10205. 50. Birdsell DN, Stewart T, Vogler AJ, Lawaczeck E, Diggs A, Sylvester TL, Buchhagen JL, Auerbach RK, Keim P, Wagner DM. 2009. Francisella tularensis subsp. novicida isolated from a human in Arizona. BMC Res Notes 2:223. 51. Clarridge JE III, Raich TJ, Sjöstedt A, Sandström G, Darouiche RO, Shawar RM, Georghiou PR, Osting C,

52.

53.

54. 55.

56.

57.

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

68.

n

861

Vo L. 1996. Characterization of two unusual clinically significant Francisella strains. J Clin Microbiol 34:1995– 2000. Birkbeck TH, Feist SW, Verner-Jeffreys DW. 2011. Francisella infections in fish and shellfish. J Fish Dis 34:173– 187. Overholt EI, Tigertt WD, Kadull PJ, Ward MK, Charkes ND, Rene RM, Salzman TE, Stephens M. 1961. An analysis of forty-two cases of laboratory-acquired tularemia. Am J Med 30:785–806. Pike RM. 1976. Laboratory-associated infections: summary and analysis of 3921 cases. Health Lab Sci 13:105–114. Shapiro DS, Schwartz DR. 2002. Exposure of laboratory workers to Francisella tularensis despite a bioterrorism procedure. J Clin Microbiol 40:2278–2281. Department of Health and Human Services, Centers for Disease Control and Prevention, and National Institutes of Health. 2007. Biosafety in Microbiological and Biomedical Laboratories, 5th ed. Department of Health and Human Services, Centers for Disease Control and Prevention and National Institutes of Health, Washington, DC. http:// www.cdc.gov/od/ohs/biosfty/bmbl5/bmbl5toc.htm. Johansson A, Berglund L, Eriksson U, Göransson I, Wollin R, Forsman M, Tärnvik A, Sjöstedt A. 2000. Comparative analysis of PCR versus culture for diagnosis of ulceroglandular tularemia. J Clin Microbiol 38:22–26. Guarner J, Breer PR, Bartlett JC, Chu MC, Shieh WJ, Zaki SR. 1999. Immunohistochemical detection of Francisella tularensis in formalin-fixed paraffin-embedded tissue. Appl Immunohistochem Mol Morphol 7:122–126. Sjöstedt A, Eriksson U, Berglund L, Tärnvik A. 1997. Detection of Francisella tularensis in ulcers of patients with tularemia by PCR. J Clin Microbiol 35:1045–1048. Celebi G, Baruönü F, Ayogˇ lu F, Cinar F, Karadenizli A, Ugˇ ur MB, Gedikogˇ lu S. 2006. Tularemia, a reemerging disease in northwest Turkey: epidemiological investigation and evaluation of treatment responses. Jpn J Infect Dis 59:229–234. Versage JL, Severin DD, Chu MC, Petersen JM. 2003. Development of a multitarget real-time TaqMan PCR assay for enhanced detection of Francisella tularensis in complex specimens. J Clin Microbiol 41:5492–5499. Willke A, Meric M, Grunow R, Sayan M, Finke EJ, Splettstösser W, Seibold E, Erdogan S, Ergonul O, Yumuk Z, Gedikoglu S. 2009. An outbreak of oropharyngeal tularaemia linked to natural spring water. J Med Microbiol 58:112–116. Maurin M, Pelloux I, Brion JP, Del Banõ JN, Picard A. 2011. Human tularemia in France, 2006–2010. Clin Infect Dis 53:133–141. Ahlinder J, Öhrman C, Svensson K, Lindgren P, Johansson A, Forsman M, Larsson P, Sjödin A. 2012. Increased knowledge of Francisella genus diversity highlights the benefits of optimised DNA-based assays. BMC Microbiol 12:220. Kugeler KJ, Gurfield N, Creek JG, Mahoney KS, Versage JL, Petersen JM. 2005. Discrimination between Francisella tularensis and Francisella-like endosymbionts when screening ticks by PCR. Appl Environ Microbiol 71:7594–7597. Sréter-Lancz Z, Széll Z, Sréter T, Márialigeti K. 2008. Detection of a novel Francisella in Dermacentor reticulatus: a need for careful evaluation of PCR-based identification of Francisella tularensis in Eurasian ticks. Vector Borne Zoonotic Dis 9:123–126. Barns SM, Grow CC, Okinaka RT, Keim P, Kuske CR. 2005. Detection of diverse new Francisella-like bacteria in environmental samples. Appl Environ Microbiol 71:5494– 5500. Petersen JM, Schriefer ME, Gage KL, Montenieri JA, Carter LG, Stanley M, Chu MC. 2004. Methods for

862 n

69.

70.

71.

72.

73.

74.

75.

76.

77. 78.

79.

80.

81.

82.

83.

84.

BACTERIOLOGY

enhanced culture recovery of Francisella tularensis. Appl Environ Microbiol 70:3733–3735. Thomas RM, Titball RW, Oyston PC, Griffin K, Waters E, Hitchen PG, Michell SL, Grice ID, Wilson JC, Prior JL. 2007. The immunologically distinct O antigens from Francisella tularensis subspecies tularensis and Francisella novicida are both virulence determinants and protective antigens. Infect Immun 75:371–378. Forsman M, Sandström G, Jaurin B. 1990. Identification of Francisella species and discrimination of type A and type B strains of F. tularensis by 16S rRNA analysis. Appl Environ Microbiol 56:949–955. Jantzen E, Berdal BP, Omland T. 1979. Cellular fatty acid composition of Francisella tularensis. J Clin Microbiol 10:928–930. Seibold E, Maier T, Kostrzewa M, Zeman E, Splettstoesser W. 2010. Identification of Francisella tularensis by whole-cell matrix-assisted laser desorption ionization–time of flight mass spectrometry: fast, reliable, robust, and costeffective differentiation on species and subspecies levels. J Clin Microbiol 48:1061–1069. Sandström G, Sjöstedt A, Forsman M, Pavlovich NV, Mishankin BN. 1992. Characterization and classification of strains of Francisella tularensis isolated in the Central Asian focus of the Soviet Union, and in Japan. J Clin Microbiol 30:172–175. Molins CR, Carlson JK, Coombs J, Petersen JM. 2009. Identification of Francisella tularensis subsp. tularensis A1 and A2 infections by real-time polymerase chain reaction. Diagn Microbiol Infect Dis 64:6–12. Johansson A, Farlow J, Larsson P, Dukerich M, Chambers E, Byström M, Fox J, Chu M, Forsman M, Sjöstedt A, Keim P. 2004. Worldwide genetic relationships among Francisella tularensis isolates determined by multiple-locus variable-number tandem repeat analysis. J Bacteriol 186:5808–5818. Vogler AJ, Birdsell D, Wagner DM, Keim P. 2009. An optimized, multiplexed multi-locus variable-number tandem repeat analysis system for genotyping Francisella tularensis. Lett Appl Microbiol 48:140–144. Tärnvik A. 1989. Nature of protective immunity to Francisella tularensis. Rev Infect Dis 11:440–451. Viljanen MK, Nurmi T, Salminen A. 1983. Enzymelinked immunosorbent assay (ELISA) with bacterial sonicate antigen for IgM, IgA, and IgG antibodies to Francisella tularensis: comparison with bacterial agglutination test and ELISA with lipopolysaccharide antigen. J Infect Dis 148:715–720. Bevanger L, Maeland JA, Kvam AI. 1994. Comparative analysis of antibodies to Francisella tularensis antigens during the acute phase of tularemia and eight years later. Clin Diagn Lab Immunol 1:238–240. Waag DM, McKee KT Jr, Sandström G, Pratt LLK, Bolt CR, England MJ, Nelson GO, Williams JC. 1995. Cellmediated and humoral immune responses after vaccination of human volunteers with the live vaccine strain of Francisella tularensis. Clin Diagn Lab Immunol 2:143–148. Brown SL, McKinney FT, Klein GC, Jones WL. 1980. Evaluation of a safranin-O-stained antigen microagglutination test for Francisella tularensis antibodies. J Clin Microbiol 11:146–148. Behan KA, Kleinn GC. 1982. Reduction of Brucella species and Francisella tularensis cross-reacting agglutinins by dithiothreitol. J Clin Microbiol 16:756–757. Ohara S, Sato T, Homma M. 1974. Serological studies on Francisella tularensis, Francisella novicida, Yersinia philomiragia, and Brucella abortus. Int J Syst Bacteriol 24:191–196. Porsch-Özcürümez M, Kischel N, Priebe H, Splettstösser W, Finke EJ, Grunow R. 2004. Comparison of enzyme-

85.

86.

87.

88.

89.

90.

91.

92.

93.

94.

95.

96.

97.

98.

99.

100.

linked immunosorbent assay, Western blotting, microagglutination, indirect immuno-fluorescence assay, and flow cytometric serological diagnosis of tularemia. Clin Diagn Lab Immunol 11:1008–1015. Schmitt P, Splettstosser W, Porsch-Özcürümez M, Finke EJ, Grunow R. 2005. A novel screening ELISA and a confirmatory Western blot useful for diagnosis and epidemiological studies of tularemia. Epidemiol Infect 133:759–766. Ericsson M, Sandström G, Sjöstedt A, Tärnvik A. 1994. Persistence of cell-mediated immunity and decline of humoral immunity to the intracellular bacterium Francisella tularensis. J Infect Dis 170:110–114. Surcel HM, Ilonen J, Poikonen K, Herva E. 1989. Francisella tularensis-specific T-cell clones are human leukocyte antigen class II restricted, secrete interleukin-2 and gamma interferon, and induce immunoglobulin production. Infect Immun 57:2906–2908. Surcel HM, Sarvas M, Helander IM, Herva E. 1989. Membrane proteins of Francisella tularensis LVS differ in ability to induce proliferation of lymphocytes from tularemia-vaccinated individuals. Microb Pathog 7:411–419. Eliasson H, Olcén P, Sjöstedt A, Jurstrand M, Bäck E, Andersson S. 2008. Kinetics of the immune response associated with tularemia: comparison of an enzymelinked immunosorbent assay, a tube agglutination test, and a novel whole-blood lymphocyte stimulation test. Clin Vaccine Immunol 15:1238–1243. Tärnvik A, Chu MC. 2007. New approaches to diagnosis and therapy of tularemia. Ann N Y Acad Sci 1105:378– 404. Urich SK, Petersen JM. 2008. In vitro susceptibility of isolates of Francisella tularensis types A and B from North America. Antimicrob Agents Chemother 52:2276–2278. Georgi E, Schacht E, Scholz HC, Splettstoesser WD. 2012. Standardized broth microdilution antimicrobial susceptibility testing of Francisella tularensis subsp. holarctica strains from Europe and rare Francisella species. J Antimicrob Chemother 67:2429–2433. Kaya A, Uysal IÖ, Güven AS, Engin A, Gültürk A, I˙çagˇ asıogˇ lu FD, Cevit Ö. 2011. Treatment failure of gentamicin in pediatric patients with oropharyngeal tularemia. Med Sci Monit 17:CR376–CR380. Meric M, Willke A, Finke EJ, Grunow R, Sayan M, Erdogan S, Gedikoglu S. 2008. Evaluation of clinical, laboratory, and therapeutic features of 145 tularemia cases: the role of quinolones in oropharyngeal tularemia. APMIS 116:66–73. Antunes NT, Frase H, Toth M, Vakulenko SB. 2012. The class A β-lactamase FTU-1 is native to Francisella tularensis. Antimicrob Agents Chemother 56:666–671. Cross JT, Jacobs RF. 1993. Tularemia: treatment failures with outpatient use of ceftriaxone. Clin Infect Dis 17:976– 980. Kudelina R I, Olsufiev NG. 1980. Sensitivity to macrolide antibiotics and lincomycin in Francisella tularensis holarctica. J Hyg Epidemiol Microbiol Immunol 24:84–91. Clinical and Laboratory Standards Institute. 2006. Methods for Dilution Antimicrobial Susceptibility Tests for Bacteria That Grow Aerobically. CLSI Document M7-A7. Approved Standard, 7th ed. Clinical and Laboratory Standards Institute, Wayne, PA. Clinical and Laboratory Standards Institute. 2009. Performance Standards for Antimicrobial Susceptibility Testing; Nineteenth Informational Supplement. M100-S19. Clinical and Laboratory Standards Institute, Wayne, PA. Federal Register. 2012. Possession, use, and transfer of select agents and toxins; biennial review; final rule. 42 CFR Part 73. Fed Regist 77:61083–61115.

Brucella* GEORGE F. ARAJ

47 Brucella spp. are common zoonoses among domestic animals and among wildlife, including novel species of marine mammals. Brucella spp. also cause infections in humans and can mimic other infectious and noninfectious diseases, posing challenges to physicians in reaching a diagnosis. The remittent/undulant fever of brucellosis was first confused with other diseases, such as malaria and typhoid fever, and has had many synonyms pertaining mainly to the geographic locations where the disease occurred: Mediterranean fever, Malta fever, Gibraltar fever, and Cyprus fever (1, 2). Over the last decade, there has been renewed interest in this organism due to its inclusion in the potential biological weapons lists of most authorities (3, 4).

cally from the six terrestrial species by their patterns of substrate-mediated metabolic activity. Though preferred or predominant hosts are recognized for Brucella spp., crossinfection of other mammalian species, including humans, may occur (12).

DESCRIPTION OF THE GENUS Brucella spp. are facultative, intracellular, small (0.5- to 1.5-mm), Gram-negative coccobacilli that lack capsules, flagella, endospores, or native plasmids. They are aerobic (some prefer CO2 for their growth), do not ferment sugars, and are positive in a few oxidative metabolic tests. Brucella spp. can grow on a wide range of culture media, and colonies appear after 24 to 48 h of incubation as mostly smooth colonies, but rough variants can occur (12, 13).

TAXONOMY AND GENOME Brucellaceae is a family of phylogenetically closely related free-living soil organisms composed of Brucella, Ochrobactrum, and Mycoplana spp. The Brucellaceae are part of the order Rhizobiales, which includes other genera involved in human disease: Bartonella, Afipia, Methylobacterium, and Roseomonas (5, 6). The taxonomy of Brucella spp. remains to be clarified. Studies indicate that terrestrial Brucella spp. are homogeneous species harboring >90% interspecies homology by DNADNA hybridization studies, identical 16S rRNA gene sequences, and >98% sequence homology by comparative genomics. Because of these findings, a suggestion was made to consider Brucella a monospecific genus and the different species as biovars of Brucella melitensis (7). The average size of the genome is 2.37 × 109 Da, with a DNA G +C content of 58 to 59 mol%. Currently, the genus Brucella encompasses 10 recognized species: 6 terrestrial, 3 marine, and 1 proposed species of unknown origin (Brucella inopinata sp. nov.) from a breast implant (8–11). The six terrestrial Brucella species are B. melitensis (three biovars) (preferred hosts are goats, sheep, and camels), B. abortus (seven biovars) (cattle and buffaloes), B. suis (five biovars) (swine and a range of wild animals), B. canis (dogs), B. ovis (rams), and B. neotomae (desert and wood rats). The three identified marine species, B. delphini, B. pinnipedialis, and B. ceti, were recovered from marine mammals (e.g. seals, whales, and dolphins) and were found to differ phenotypi-

ANTIGENIC COMPONENTS Several antigenic determinants of Brucella, related mainly to lipopolysaccharide (LPS) and protein antigens, have been characterized. The LPS is the major antigen that dominates the antibody response. LPS of rough strains is very similar to LPS of smooth strains. Based on their O side chain, smooth strains were reported to be composed of two antigenic epitopes: A (B. abortus) and M (B. melitensis). The smooth-strain LPS has been reported to be responsible for observed cross-reactions in both the agglutination and complement fixation tests between smooth species of Brucella and Yersinia enterocolitica O:9, Escherichia hermannii, Escherichia coli O:157, Salmonella enterica serovar O:30, Stenotrophomonas maltophilia, Vibrio cholerae O:1, and Francisella tularensis. Cross-reaction has been attributed to the similarities of the O-specific side chains of the LPS molecules of these organisms (reference 14; refer also to chapter 46 in this Manual). The characterized protein antigens include outer and inner membrane, cytoplasmic, and periplasmic antigens. Some are recognized by the immune system during infection and are potentially useful in diagnostic tests (15). For example, Omp25 is an outer membrane structural protein that is highly conserved in all brucellae and is associated with both LPS and peptidoglycan. In addition, some proteins, such as ribosomal proteins (e.g., L7/L12) and fusion proteins, demonstrate a protective effect against Brucella based on antibody and cell-mediated responses (16). These molecules may be useful in potential vaccines.

*This chapter contains information presented in chapter 44 by Jeannine M. Petersen, Martin E. Schriefer, and George F. Araj in the 10th edition of this Manual.

doi:10.1128/9781555817381.ch47

863

864 n

BACTERIOLOGY

VIRULENCE FACTORS, PATHOGENIC MECHANISMS, AND IMMUNE RESPONSE The exact pathophysiologic aspects of infection remain to be defined, especially since this “stealth pathogen” can infect and survive without inducing a massive inflammatory response (17–20). The incubation period is variable but generally is 1 to 4 weeks. The intracellular location and survival of the organism contribute to its virulence and pathogenesis. Virulence determinants include urease to avoid stomach stress through oral passage and Brucella-containing vacuoles (BCV), which enable escape from immune system recognition and provide an acidic environment to hamper antibiotic activity. A Brucella LPS cell component (containing a poly N-formyl perosamine O chain and a CuZn superoxide dismutase) and outer membrane protein 25 (OMP25) help the bacteria survive within mononuclear phagocytes (21). Also, Brucella LPS poorly induces gamma interferon and tumor necrosis factor alpha (TNF-α), both of which are essential for T-helper 1 (Th1)-type-cell-mediated immunity for the elimination of the organism (22). Briefly, once the brucellae enter the body by various routes, they are encountered by polymorphonuclear and mononuclear phagocytes, to which lectin facilitates attachment, and lipid rafts help in avoidance of defense mechanisms. After entry into the cell, the first step entails the establishment of an intracellular replication niche, and Brucella LPS and a periplasmic cyclic β-glucan are essential for this step. Brucella spp. escape from phagocytic killing by inhibiting the phagosome-lysosome fusion and hence evade the immune system. These factors together with the BCV aid in its survival, propagation, and persistence within macrophages and other cells. As brucellae remain within macrophages, they are transported through regional lymph nodes into the circulatory system and subsequently seeded throughout the body, with tropism for the reticuloendothelial system, resulting in different clinical phases of disease (18). The overall inflammatory process results in a slow degradation of Brucella cell wall components by the polymorphonuclear leukocytes and can lead to granuloma formation, which is more often associated with B. abortus than B. melitensis (18). Protective immunity, though not long term, is conferred by antibodies to LPS and T-cell-mediated macrophage activation, triggered by protein antigens. Lymphocytes are the main stimulant of the immune response. The Th1 response stimulates IgG2a production, which is involved mostly in protection against intracellular pathogens through cell-mediated immunity, and is critical for the clearance of Brucella infection. The Th2 response stimulates the production of IgG1 and is mainly responsible for protection against extracellular pathogens through the humoral immune response (23, 24). Recently, the production of cytokines, chemokines, and matrix-metalloproteinases has been associated with induced osteoclastogenesis in brucella arthritis and osteomyelitis, and the OMP19 lipoprotein, together with TNF-α, were reported to be associated with astrocyte apoptosis in neurobrucellosis (23). Though receiving close attention, the exact nature of the immune response and the protective antigens/ factors involved in this disease are still being investigated, and the pathogenic mechanisms of reinfection remain unknown (25).

EPIDEMIOLOGY AND TRANSMISSION Animals are generally asymptomatic carriers of Brucella spp. The major symptoms appear during infectious abortion of

the animal placenta and fetus. Invaded fetal tissue may contain up to 1010 infectious bacteria per gram of tissue and fluid (12). Although Brucella can be killed by pasteurization, exposure to UV light, acidity, or many antiseptics and disinfectants, it can survive for long periods under various conditions, e.g., 10 weeks in soil, 11 weeks in aborted fetuses, 17 weeks in bovine stool, around 3 weeks in milk and ice cream, and several months in fresh goat cheese (12, 26). Brucella can infect both humans and animals. In terms of the total numbers of infected human cases, B. melitensis dominates the world arena (especially in the Mediterranean and Arabian Gulf countries). However, B. abortus and B. suis supersede it in certain geographic locations in Europe. B. canis has also been reported to cause human diseases, while B. ovis and B. neotomae have not (12, 27–29). Brucella spp. associated with marine animals have been reported to cause disease in humans (30, 31). The epidemiologies of human brucellosis differ between areas of endemicity and nonendemicity in terms of age, sex, season, and risk factors. In regions of endemicity, such as the eastern Mediterranean Basin, Middle East, the Arabian Peninsula, Mexico, Central and South America, Southern Europe, Central Asia, and the Indian subcontinent, the disease occurs among the general population. In the general population, levels of infection are almost equal among adults and children of both sexes and mostly due to ingestion of unpasteurized goat, sheep, cow, and camel milk or its products (e.g., soft cheese, ice cream) (2, 27–29, 32–34). In areas where the disease is not endemic, infection is seen predominantly among adult males, acquired occupationally by transmission through direct skin contact (e.g., through cuts and abrasions) with infected animal parts, inhalation of aerosolized infected particles, and accidental inoculation (e.g., sprays or aerosols inoculated into the eye, mouth, and nose). These infections occur mostly among dairy industry professionals, veterinarians, abattoir workers, and clinical and research microbiology staff (35, 36). Human-to-human transmission may occur, although very rarely. Few cases of neonatal brucellosis have been reported, and the isolation of Brucella from human milk may explain its pathogenesis (37, 38). Laboratory-acquired infection (LAI) is an important source of transmission. Brucella has a very low infectious dose (≤102 organisms), and personnel should adhere to strict safety precautions, especially when handling cultures suspected of containing the organisms in clinical, research, and production laboratories (36, 39, 40). Most cases of laboratory-acquired disease result from mishandling and misidentification of the organism (40). The frequent failure of clinical laboratories to correctly identify isolates as Brucella species is particularly worrisome from the perspective of laboratory safety and potential use as a bioweapon. B. melitensis, B. abortus, and B. suis are category B select agents (4).

CLINICAL CATEGORIES OF HUMAN BRUCELLOSIS The clinical categories of human brucellosis are based on arbitrary criteria. In 1956, Spink based them on the duration of symptoms (acute, 12 months) (41). Subsequently, others based them primarily on extent of clinical manifestations (e.g., subclinical, localized, chronic, and active, with or without localized disease, including bacteremic and serological classifications) (29, 42). To date, no uniform definition has been adopted.

47. Brucella

The incubation period is variable but usually ranges between 1 and 4 weeks. The disease onset is insidious and can present with a wide and diverse range of nonspecific clinical signs and symptoms, such as fever, sweats, arthralgias, myalgia, fatigue, loss of appetite, weight loss, hepatomegaly, and splenomegaly. Complications can involve many organs and tissues with signs of focal disease. The routine hematology and biochemical profiles are usually within normal limits, with some elevation in erythrocyte sedimentation rate and liver function tests. Thus, to the unaware physician, the diagnosis of brucellosis can be a dilemma and may protract for weeks and, in some complicated cases, for years (27, 29, 43–47). Increased business and leisure travel to countries where the disease is endemic has led to diagnostic challenges in areas where brucellosis is uncommon, especially when the presentation is unusual (29, 48). Overall, the mortality is very low, but morbidity is high. Previously thought to be uncommon, brucellosis in childhood now seems to be as prevalent as and presents in a manner similar to that in adults in areas of endemicity (33, 49). Because of these nonspecific clinical features, human brucellosis can imitate a variety of diseases and, thus, has been labeled “the disease of mistakes.” For example, it can be misdiagnosed and confused with other diseases, such as typhoid fever, rheumatic fever, tuberculosis, malaria, infectious mononucleosis, endocarditis, histoplasmosis, ankylosing spondylitis, pyelitis, cholecystitis, thrombophlebitis, chronic fatigue syndrome, collagen vascular diseases, autoimmune diseases, and tumors (27, 29, 46, 47, 50).

COMPLICATIONS The most commonly encountered focal complications are osteoarticular (10 to 70%) (mostly joints), genital in both males (6 to 8%) and females (2 to 5%), neurological (3 to 5%), cardiac (1 to 3%), pulmonary (1 to 2%), and renal (150 μm Taenia spp. (T. saginata, beef tapeworm; T. solium, pork tapeworm) Hymenolepis nana (dwarf tapeworm)

Trichuris trichiura (whipworm) Enterobius vermicularis (pinworm) Ascaris lumbricoides (large roundworm), fertilized eggs Hookworm Diphyllobothrium spp, (broad fish tapeworm)

Hymenolepis diminuta (rat tapeworm)

Paragonimus spp. (lung fluke)

Trichostrongylus spp. Ascaris lumbricoides (large roundworm); unfertilized eggs Schistosoma japonicum (blood fluke, stool) Schistosoma mekongi (rounder and smaller than S. japonicum); measure 50–65 by 30–55 μm Schistosoma haematobium (blood fluke, urine) Schistosoma intercalatum (blood fluke, stool) Schistosoma mansoni (blood fluke, stool) Fasciolopsis buski (giant intestinal fluke) or Fasciola hepatica (sheep liver fluke) or Echinostoma spp.

a

This table does not include every possible helminth that could be found as a human parasite; however, the most likely helminth infections are included.

histolytica-Entamoeba dispar group, and Entamoeba histolytica (Table 6). These methods (enzyme immunoassay [EIA], fluorescent-antibody assay [FA], and immunochromatographic assay [cartridge]) are designed to detect the antigens of select organisms; a negative result does not rule out the possibility that these organisms/antigen are present in low numbers or that other intestinal parasites are etiologic agents causing disease, including Dientamoeba fragilis, the microsporidia, and helminth parasites (1, 17–19). Immunoassay reagents are currently under development and trial for D.

MCM 11th Edition X : MCM11$C135 03-22-15 03:32:47

Page 2320

fragilis and Blastocystis spp. Fecal immunoassays for the microsporidia are available; however, none are currently FDAapproved for use within the United States.

Molecular Methods In January 2013, the FDA approved the first test that can simultaneously detect 11 common viral, bacterial, and parasitic (Cryptosporidium, Giardia) causes of infectious gastroenteritis from a single patient sample (xTAG Gastrointestinal Pathogen Panel; Luminex, Inc., Austin, TX)

135. General Approaches for Detection and Identification of Parasites n 2321 TABLE 3

Diagnostic characteristics of organisms in permanent stained smears

Specimen

Protozoa

Stool, other specimens from gastrointestinal tract, urogenital system

Size, shape, stage (trophozoite, precyst, cyst, oocyst, spore) Nuclear arrangement, cytoplasm inclusions (chromatoidal bars, vacuoles, axonemes, axostyles, median bodies, sporozoites, polar tubules) Balantidium coli trophozoites and cysts may not be visible due to excess stain retention.

There are also several molecular tests that are in clinical trials for the detection of select gastrointestinal parasites. These tests are molecular gastrointestinal panels and target the most commonly occurring bacterial, viral, and parasitic stool pathogens. Although there are laboratory-developed tests for most parasites, these are not commercially available or available only in specialized testing centers. When such tests are used, there should be attention given to the use of internal amplification controls to detect inhibition, since common specimens, such as blood and stool, contain PCR inhibitors. Thorough validation is required before these are implemented for clinical testing.

ADDITIONAL TECHNIQUES FOR STOOL EXAMINATION Although the routine O&P examination consisting of the direct wet mount, the concentration, and the permanent stained smear is an excellent procedure recommended for the detection of most intestinal parasites, several other diagnostic techniques are available for the recovery and identification of specific parasitic organisms (1, 6, 10). Most labora-

TABLE 4

Helminths Egg, larvae, and/or adults may not be identified because of excess stain retention or distortion

tories do not routinely offer all of these techniques, but many can be performed relatively simply and inexpensively. Occasionally, it is necessary to examine stool specimens for the presence of scolices and proglottids of cestodes and adult nematodes and trematodes to confirm the diagnosis and/or for species identification (Table 7). A method for the recovery of these stages is also described in this chapter.

Culture of Larval-Stage Nematodes Nematode infections giving rise to larval stages that hatch in soil or in tissues may be diagnosed by using fecal culture methods to concentrate the larvae (1, 6, 10, 15). Strongyloides stercoralis larvae are the most common larvae found in stool specimens. Depending on the fecal transit time through the intestine and the patient’s condition, rhabditiform and filariform larvae may be present. Caution must be exercised when handling larval cultures because infective filariform larvae may be present. If there is a delay in the preservation of the stool specimen, then embryonated ova as well as larvae of hookworm may be present. Culture of feces for larvae is useful for (i) revealing their presence when they are too scanty to be detected by concentration methods; (ii) distinguishing whether the infection is due to

Key to identification of intestinal amebae (permanent stained smear)

1. Trophozoites present ............................................................................................................................................... 2 Cysts present ........................................................................................................................................................... 7 2. Trophozoites measure >12 μm................................................................................................................................ 3 Trophozoites measure 10 μm (including any shrinkage “halo”)..................................................................................... 8 Cysts measure 10 μm ........... Cytostome present, 10 μm ........................... Two nuclei, no fibrils, 80%) than the wet mount method and is considered the gold standard method for the detection of T. vaginalis. Specimens must be collected properly and inoculated immediately into the appropriate medium, such as modified Diamond’s, Trichosel, or Hollander’s medium. Due to cost and convenience, this approach is not routinely used. Culture systems (InPouchTV [BioMed Diagnostics, San Jose, CA] and the system of Empyrean Diagnostics, Inc. [Mountain View, CA]) that allow direct inoculation, transport, culture, and microscopic examination are commercially available (166, 167). In situations in which immediate transport of specimens is not

140. Intestinal and Urogenital Parasites n 2415

feasible, the use of these transport/culture devices should be encouraged. Studies have also shown that a delayed inoculation protocol is as sensitive as immediate inoculation, allowing the results of microscopy to be used to determine whether further culture is necessary (168). Serologic testing is not useful for the diagnosis of trichomoniasis.

Antigen Detection Several antigen detection methods have been developed for T. vaginalis and offer the advantage of being rapid and easy to perform. A latex agglutination test (TV Latex; Kalon Biological, Guildford, United Kingdom) has been shown to have excellent sensitivity (169) but is not available in the United States. An immunofluorescence assay (Light Diagnostics T. vaginalis DFA; Chemicon International, Temecula, CA) is available in the United States for testing directly from patient samples. An immunochromatographic capillary flow assay is available commercially for the qualitative detection of T. vaginalis antigens from vaginal swabs. The OSOM Trichomonas Rapid Test Kit (Sekisui Diagnostics, Tokyo, Japan) is a dipstick assay providing results in 10 min. In published studies, the OSOM test has demonstrated good sensitivity (67 to 94.7%) and specificity (98.8 to 100%) compared to various comparator assays, including wet mount, culture, and amplified testing (170–176). This assay would perhaps be more suitable for a clinic setting, where more-rapid results are desirable. A rapid point-ofcare test is being developed for T. vaginalis. This assay features novel electrochemical endpoint detection and in initial studies appears to have high sensitivity and specificity (177).

Nucleic Acid Detection Techniques The Affirm VPIII (Becton Dickinson) is a direct DNA probe test for the detection of organisms from vaginal swabs associated with vaginosis/vaginitis. It tests for the three most common syndromes associated with increased vaginal discharge: bacterial vaginosis (Gardnerella vaginalis), candidiasis (Candida albicans), and trichomoniasis (T. vaginalis). According to the manufacturer’s package insert, the assay has a sensitivity and specificity of 90 and 98%, respectively, compared with wet mount and culture for T. vaginalis (package insert, Affirm VPIII, version no. 670160JAAG; Becton Dickinson). In a clinical evaluation of vaginal swab specimens from both symptomatic and asymptomatic females, the Affirm detected more T. vaginalis-positive samples than wet mount testing, although the difference was not statistically significant (178). Nucleic acid-based amplification methods, such as PCR and transcription-mediated amplification (TMA), developed for the detection of T. vaginalis have been reported in the literature, though only one is currently FDA cleared. These amplification methods have demonstrated varying sensitivities depending on the genomic target, specimen type, and sex of the patient (172, 179–183). PCR has a sensitivity of 85 to 100% with vaginal swabs; its sensitivity with urine is lower, ranging from 60 to 80% (153, 184, 185). For men, PCR with urine and urethral swabs has been reported and appears to be more sensitive than conventional methods (186, 187). Several researchers have reported using TMA technology from Hologic/Gen-Probe (San Diego, CA) for detection of T. vaginalis, which is now FDA approved (148, 181, 182, 188, 189). Studies using the Trichomonas Aptima Combo 2 assay have shown the performance of the assay to be superior compared to other methodologies (147, 148, 189). The assay is approved for the detection of T. vaginalis infections from a wide variety of specimens

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2415

such as clinician-collected vaginal specimens, endocervical swabs, ThinPrep liquid-based cytology samples, and urine samples (189). Samples can be automated for testing using either the Panther or the Tigris instrumentation. In addition to testing on adults, molecular testing is also being evaluated for the detection of Trichomonas in children (190). Some researchers have evaluated the Hologic/Gen-Probe assay for diagnosis of anorectal T. vaginalis using rectal swabs (146, 191). Another assay available outside the United States is the Seeplex STD 6 ACE Detection System (Seegene, Seoul, Korea), which detects T. vaginalis and other pathogens (192). This assay is a multiplex PCR assay. Detection employs six pairs of dual priming oligonucleotide primers targeted to unique genes of the specific pathogen. It is not clear if this assay will receive approval in the United States. While the wet mount method provides a rapid result at a low cost, tests with the increased sensitivities, such as nucleic acid probes or amplification tests, may be indicated because of the impact of T. vaginalis infections on pregnancy and the link with HIV transmission (165, 187). The use of alternate specimen types such as urine makes amplified testing an important advancement for diagnosis of trichomoniasis. An algorithm to reflex specimens with negative wet mounts to culture or a more sensitive methodology may be a useful diagnostic approach (173). Because the true prevalence of T. vaginalis is unknown and the prevalence appears to be higher in older women, it is important for laboratories and clinicians to be aligned in regards to which patients should be tested (148).

Treatment The recommended treatment for T. vaginalis infections is metronidazole or tinidazole (62, 193). For metronidazole, oral therapy is recommended over topical treatment. Tinidazole may be used as a first-line agent or for refractory cases previously treated with metronidazole (194). For treatment during pregnancy, metronidazole is the recommended therapy; however, it should be used cautiously, as data do not suggest that metronidazole treatment results in a reduction in perinatal morbidity (193, 195). All sexual partners of infected individuals should also receive treatment. Treatment failure with metronidazole is most often due to noncompliance or reinfection. True resistance to metronidazole has been documented and appears to be increasing (194, 196). While not routinely available, methods have been published for the in vitro determination of susceptibility. These methods have not been standardized, and the results can vary based on assay conditions (197, 198).

Evaluation, Interpretation, and Reporting of Results A laboratory finding that is positive for T. vaginalis is considered diagnostic of trichomoniasis. As discussed above, both microscopy and culture are prone to lower sensitivities due to issues related to sampling and transport. Laboratories should have strict rejection criteria for Trichomonas culture and wet mount specimens that do not arrive within the specified time or transport conditions; such policies improve sensitivity, ensuring more-accurate results. In comparison to methods such as antigen detection and molecular methods, a negative result by these methods should be viewed cautiously and evaluated in conjunction with clinical symptoms. With testing for Trichomonas, such as antigen detection and direct molecular probes or amplified tests, reported results should reflect the analyte that is detected. For example, a positive result for T. vaginalis by a direct DNA probe assay should state “T. vaginalis DNA detected.” If testing is

2416 n PARASITOLOGY

expanded to differing sample types, such as urine testing by amplified methods, the report should clearly state the specimen tested. If a home-brew assay is used for the diagnosis of Trichomonas, it is required that the result be labeled to indicate its status as an in-house test in accordance with Clinical Laboratory Improvement Amendments regulations as follows: “This test was developed and its performance characteristics determined by [Laboratory Name]. It has not been cleared or approved by the U.S. Food and Drug Administration.”

specimens are examined, it is important to note that cysts of E. hominis can resemble those of E. nana, although E. nana cysts containing two nuclei are rare. Because of the small sizes of E. hominis and R. intestinalis, it is difficult to detect these organisms even when permanent-stained smears are examined. This may lead to the underreporting of both organisms. R. intestinalis has been recovered from the pancreatic juice of a patient with small lesions of the pancreatic duct (199). In general, treatment is not recommended for infections with the nonpathogenic flagellates. Improved personal hygiene and sanitary conditions are key methods for the prevention of infection.

NONPATHOGENIC FLAGELLATES C. mesnili is found worldwide and is generally considered nonpathogenic. Unlike D. fragilis, C. mesnili has both a trophozoite and a cyst stage. The organism is acquired through the ingestion of contaminated food or water and resides in the cecum and/or colon of the infected human or animal. The trophozoite is 6 to 24 μm long and contains a characteristic spiral groove that runs longitudinally along the body (Table 4; Fig. 7 and 11). Motility of the organism can sometimes be seen in fresh preparations, and the spiral groove may be exposed as the organism turns. Flagella are difficult to see in stained preparations. The trophozoite contains one nucleus, with a cytostome or oral groove in close proximity. The pear-shaped cyst retains the cytoplasmic organelles of the trophozoite, with a single nucleus and curved cytostomal fibril. Observing the organism in permanent-stained preparations makes identification more definitive. P. hominis, formerly referred to as Trichomonas hominis, is a nonpathogenic flagellate that is similar to D. fragilis in that only the trophozoite stage has been observed. Although the organism is cosmopolitan in nature and is recovered from individuals with diarrhea, it is still considered nonpathogenic. The trophozoites typically inhabit the cecum. They are pyriform and contain an undulating membrane that runs the length of the parasite. The use of permanent smears is recommended for observation of these organisms in clinical specimens. The trophozoites may stain weakly, making them difficult to detect on stained smears (3). Two additional nonpathogenic intestinal flagellates are E. hominis and R. intestinalis. Both E. hominis and R. intestinalis are found in warm or temperate climates, and infection is acquired through the ingestion of cysts. When clinical

CILIATES Balantidium coli Taxonomy B. coli is a ciliate belonging to the phylum Ciliophora, class Litostomatea, order Trichostomatia (see chapter 132 of this Manual). Members of the phylum Ciliophora are protozoa possessing cilia in at least one stage of their life cycles. They also have two different types of nuclei, one macronucleus and one or more micronuclei. Over the last several years, molecular analysis has aided in the characterization of the genus Balantidium (200–202). Sequences of the genus are on file at GenBank based on the small-subunit rRNA. There is question as to whether the species isolated from humans, B. coli, and pigs, B. suis, are the same species (200).

Description of the Agent This organism has both the trophozoite and cyst forms as part of its life cycle (Table 5; Fig. 12). The cyst form is the infective stage. After ingestion of the cysts and excystation, trophozoites secrete hyaluronidase, which aids in the invasion of the tissue. The trophozoite, which is oval and covered with cilia, is easily seen in wet mount preparations under

TABLE 5 Key features of the ciliate B. coli Stage

Cyst

FIGURE 11 Chilomastix mesnili trophozoite (left) and cyst (right). Note the oral groove (feeding groove) at the right side of the trophozoite (clear area) and the curved fibril (“shepherd’s crook”) in the cyst. Also note the typical pear or lemon shape of the cyst. Organisms are stained with Wheatley’s trichrome stain. Courtesy of L. Garcia. doi:10.1128/9781555817381.ch140.f11

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2416

Characteristics

Trophozoite Shape and size: Ovoid with tapering anterior end; 50–100 μm long, 40–70 μm wide Motility: Rotary, boring; may be rapid Nuclei: 1 large kidney-bean-shaped macronucleus may not be distinct in unstained preparation; 1 small round micronucleus adjacent to macronucleus, difficult to see Cytoplasm: May be vacuolated; may contain ingested bacteria and debris; anterior cytostome Cilia: Body surface covered with longitudinal rows of cilia; longer near cytostome Note: May be confused with helminth eggs or debris on a permanent-stained smear; concentration or sedimentation examination recommended Shape and size: Spherical or oval; 50–70 μm in diam Nuclei: 1 large macronucleus, 1 micronucleus, difficult to see Cytoplasm: Vacuoles are visible in young cysts; in older cysts, internal structure appears granular Cilia: Difficult to see within the thick cyst wall

140. Intestinal and Urogenital Parasites n 2417

occurred in patients who are either elderly or immunocompromised. The disease presents as a pneumonia-like illness. In these documented cases, Balantidium has been recovered from specimens such as bronchial secretions and bronchial lavage specimens. It is hypothesized that extraintestinal colonization can occur between the lymphatic or circulatory system, by perforation through the colon, or through aspiration of fluid from the oral cavity (200). One case describes an individual with vertebral osteomyelitis and myelopathy, which is the first documented case of infection in the bone (210). FIGURE 12 Balantidium coli trophozoite (left) and cyst (right). Note the single macronucleus visible in both the trophozoite and the cyst. The much smaller micronucleus is not visible. Organisms are stained with Wheatley’s trichrome stain. Courtesy of L. Garcia. doi:10.1128/9781555817381.ch140.f12

low-power magnification. The cytoplasm contains both a macronucleus and a micronucleus, in addition to two contractile vacuoles. Motile trophozoites can be observed in fresh wet preparations, but the specimen must be observed soon after collection. The trophozoite is somewhat pear shaped and also contains vacuoles that may harbor debris such as cell fragments and ingested bacteria. Cyst formation takes place as the trophozoite moves down the large intestine.

Direct Examination Microscopy Either ova and parasite examination of feces or histological examination of intestinal biopsy specimens establishes the diagnosis of B. coli infections. The diagnosis can be established only by demonstrating the presence of trophozoites in stool or tissue samples (209). It is very easy to identify these organisms in wet preparations and concentrated stool samples. Conversely, it can be challenging to identify B. coli from trichrome-stained permanent smears because the organisms are so large and have a tendency to overstain. This makes the organism less discernible and increases the chance of misidentification.

Treatment Epidemiology, Transmission, and Prevention B. coli exists in animal reservoirs such as pigs and chimpanzees, with pigs being the primary reservoir (200). The organism is the only pathogenic ciliate and the largest pathogenic protozoan known to infect humans. Transmission occurs by the fecal-oral route following ingestion of the cysts in contaminated food or water. Infection is more common in warmer climates and in areas where humans are in close contact with pigs. As with other intestinal protozoa, poor sanitary conditions lead to a higher incidence of infection. Prevalence of Balantidium varies by geographic location but overall is estimated to be between 0.02 and 1% (203). Highprevalence areas include areas of the Middle East, Papua New Guinea and West Irian, Latin America, and the Philippines (204, 205).

The treatment of choice for B. coli infection is tetracycline, although it is considered an investigational drug when used in this context (62). Metronidazole and iodoquinol are therapeutic alternatives used in some cases (3). Nitazoxanide, which is a broad-spectrum antiparasitic drug, may be another alternative for treatment (200).

Evaluation, Interpretation, and Reporting of Results The recovery of B. coli in humans is fairly uncommon despite its worldwide distribution. Pulmonary infections can occur, but the clinical laboratory scientist needs to make sure that this organism is not confused with motile ciliated epithelial cells that can be present in respiratory specimens. Balantidium spp. in wet mounts are very active parasites with uniform ciliation.

Clinical Significance

SUMMARY Infection with B. coli is most often asymptomatic; however, symptomatic infection can occur, resulting in bouts of dysentery similar to amebiasis (3, 206). Infection with Balantidium can be described in three ways: (i) asymptomatic host, carrying the disease; (ii) chronic infection, nonbloody diarrhea, including other symptoms such as cramping and abdominal pain; and (iii) fulminating disease consisting of mucoid and bloody stools (200). In addition, colitis caused by B. coli is often indistinguishable from that caused by E. histolytica. Symptoms typically include diarrhea, nausea, vomiting, headache, and anorexia. Fluid loss can be dramatic, as seen in some patients with cryptosporidiosis. The organism can invade the submucosa of the large bowel, and ulcerative abscesses and hemorrhagic lesions can occur. The shallow ulcers and submucosal lesions that result from invasion are prone to secondary infection by bacteria and can be problematic for the patient (207, 208). Death due to invasive B. coli infection has been reported (207). Infections associated with extraintestinal sites have been described (207–209). There have been several reports of Balantidium spreading from the intestine to the lung. Most of these cases have

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2417

Clinical laboratories are now given more choices for testing in diagnostic parasitology, with assays ranging from microscopy, culture, antigen detection, and nucleic acid amplification techniques for detection of the intestinal and urogenital amebae, flagellates, and ciliates. Molecular biology has the promise to deliver more sensitive and specific methods, with the availability of an FDA-cleared assay for detection of T. vaginalis from genital specimens and a multiplex assay for E. histolytica, G. duodenalis, and Cryptosporidium species detection from stool. There are other assays being designed and in clinical trials at the time of writing, and while these methods have not been fully adapted to the clinical diagnostic laboratory, their use in the coming years is certain to increase. In addition to these amplification tests, rapid point-of-care tests are available for organisms such as T. vaginalis. Results can now be available in real time for the clinician to manage patients directly in the exam setting. While these new test modalities are exciting, clinical laboratories are still faced with using microscopy for routine workup for stool specimens due to the lack of commercially

2418 n

PARASITOLOGY

available testing for all the relevant organisms. Some laboratories have switched to antigen-based methods, but many still rely on microscopy because antigen-based methods cannot detect all potential pathogens in a given stool specimen. Microscopy, as we know, cannot differentiate between pathogenic and nonpathogenic amebae and the different genotypes of Giardia. Together these exciting new areas will help increase the options that the clinical parasitology laboratory has for the diagnosis of intestinal parasitic infections.

16.

17.

18.

REFERENCES 1. Mengelle C, Mansuy JM, Prere MF, Grouteau E, Claudet I, Kamar N, Huynh A, Plat G, Benard M, Marty N, Valentin A, Berry A, Izopet J. 2013. Simultaneous detection of gastrointestinal pathogens with a multiplex Luminex-based molecular assay in stool samples from diarrhoeic patients. Clin Microbiol Infect 19:E458–E465. 2. Clinical and Laboratory Standards Institute. 2005. Procedures for the Recovery and Identification of Parasites from the Intestinal Tract; Approved Guideline—2nd Edition. CLSI Document M28-A2. Clinical and Laboratory Standards Institute, Wayne, PA. 3. Garcia LS. 2007. Diagnostic Medical Parasitology, 5th ed. ASM Press, Washington, DC. 4. Sargeaunt PG, Williams JE. 1978. Electrophoretic isoenzyme patterns of Entamoeba histolytica and Entamoeba coli. Trans R Soc Trop Med Hyg 72:164–166. 5. Ortner S, Clark CG, Binder M, Scheiner O, Wiedermann G, Duchene M. 1997. Molecular biology of the hexokinase isoenzyme pattern that distinguishes pathogenic Entamoeba histolytica from nonpathogenic Entamoeba dispar. Mol Biochem Parasitol 86:85–94. 6. Diamond LS, Clark CG. 1993. A redescription of Entamoeba histolytica Schaudinn, 1903 (Emended Walker, 1911) separating it from Entamoeba dispar Brumpt, 1925. J Eukaryot Microbiol 40:340–344. 7. Heredia RD, Fonseca JA, Lopez MC. 2012. Entamoeba moshkovskii perspectives of a new agent to be considered in the diagnosis of amebiasis. Acta Trop 123:139–145. 8. Ali IK, Hossain MB, Roy S, Ayeh-Kumi PF, Petri WA, Jr, Haque R, Clark CG. 2003. Entamoeba moshkovskii infections in children, Bangladesh. Emerg Infect Dis 9:580– 584. 9. Clark CG, Diamond LS. 1991. The Laredo strain and other ’Entamoeba histolytica-like’ amoebae are Entamoeba moshkovskii. Mol Biochem Parasitol 46:11–18. 10. Fotedar R, Stark D, Marriott D, Ellis J, Harkness J. 2008. Entamoeba moshkovskii infections in Sydney, Australia. Eur J Clin Microbiol Infect Dis 27:133–137. 11. Herbinger KH, Fleischmann E, Weber C, Perona P, Loscher T, Bretzel G. 2011. Epidemiological, clinical, and diagnostic data on intestinal infections with Entamoeba histolytica and Entamoeba dispar among returning travelers. Infection 39:527–535. 12. Shimokawa C, Kabir M, Taniuchi M, Mondal D, Kobayashi S, Ali IK, Sobuz SU, Senba M, Houpt E, Haque R, Petri WA, Jr, Hamano S. 2012. Entamoeba moshkovskii is associated with diarrhea in infants and causes diarrhea and colitis in mice. J Infect Dis 206:744–751. 13. Royer TL, Gilchrist C, Kabir M, Arju T, Ralston KS, Haque R, Clark CG, Petri WA, Jr. 2012. Entamoeba bangladeshi nov. sp., Bangladesh. Emerg Infect Dis 18:1543– 1545. 14. Fotedar R, Stark D, Beebe N, Marriott D, Ellis J, Harkness J. 2007. Laboratory diagnostic techniques for Entamoeba species. Clin Microbiol Rev 20:511–532. 15. Walderich B, Weber A, Knobloch J. 1997. Differentiation of Entamoeba histolytica and Entamoeba dispar from German

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2418

19.

20.

21. 22.

23.

24. 25.

26.

27.

28.

29.

30. 31.

32.

33.

34.

travelers and residents of endemic areas. Am J Trop Med Hyg 57:70–74. Ortega HB, Borchardt KA, Hamilton R, Ortega P, Mahood J. 1984. Enteric pathogenic protozoa in homosexual men from San Francisco. Sex Transm Dis 11:59–63. Lowther SA, Dworkin MS, Hanson DL. 2000. Entamoeba histolytica/Entamoeba dispar infections in human immunodeficiency virus–infected patients in the United States. Clin Infect Dis 30:955–959. Hung CC, Chen PJ, Hsieh SM, Wong JM, Fang CT, Chang SC, Chen MY. 1999. Invasive amoebiasis: an emerging parasitic disease in patients infected with HIV in an area endemic for amoebic infection. AIDS 13:2421– 2428. Oh MD, Lee K, Kim E, Lee S, Kim N, Choi H, Choi MH, Chai JY, Choe K. 2000. Amoebic liver abscess in HIV-infected patients. AIDS 14:1872–1873. Hung CC, Deng HY, Hsiao WH, Hsieh SM, Hsiao CF, Chen MY, Chang SC, Su KE. 2005. Invasive amebiasis as an emerging parasitic disease in patients with human immunodeficiency virus type 1 infection in Taiwan. Arch Intern Med 165:409–415. World Health Organization. 1997. Amoebiasis. Wkly Epidemiol Rec 72:97–99. Ximenez C, Moran P, Rojas L, Valadez A, Gomez A. 2009. Reassessment of the epidemiology of amebiasis: state of the art. Infect Genet Evol 9:1023–1032. Wilson M, Schantz P, Pieniazek N. 1995. Diagnosis of parasitic infections: immunologic and molecular methods, p 1159–1170. In Murray PR, Baron EJ, Pfaller MA (ed), Manual of Clinical Microbiology, 6th ed. ASM Press, Washington, DC. Bruckner DA. 1992. Amebiasis. Clin Microbiol Rev 5:356– 369. Zaki M, Clark CG. 2001. Isolation and characterization of polymorphic DNA from Entamoeba histolytica. J Clin Microbiol 39:897–905. Espinosa-Cantellano M, Martinez-Palomo A. 2000. Pathogenesis of intestinal amebiasis: from molecules to disease. Clin Microbiol Rev 13:318–331. Ralston KS, Petri WA, Jr. 2011. Tissue destruction and invasion by Entamoeba histolytica. Trends Parasitol 27:254– 263. Petri WA, Jr, Smith RD, Schlesinger PH, Murphy CF, Ravdin JI. 1987. Isolation of the galactose-binding lectin that mediates the in vitro adherence of Entamoeba histolytica. J Clin Invest 80:1238–1244. Haque R, Neville LM, Hahn P, Petri WA, Jr. 1995. Rapid diagnosis of Entamoeba infection by using Entamoeba and Entamoeba histolytica stool antigen detection kits. J Clin Microbiol 33:2558–2561. Tanyuksel M, Petri WA, Jr. 2003. Laboratory diagnosis of amebiasis. Clin Microbiol Rev 16:713–729. Tanyuksel M, Tachibana H, Petri WA. 2001. Amebiasis, an emerging disease, p 197–202. In Scheld WM, Craig WA, Hughes JM (ed), Emerging Infections 5. ASM Press, Washington, DC. Garcia LS, Shimizu RY, Bernard CN. 2000. Detection of Giardia lamblia, Entamoeba histolytica/Entamoeba dispar, and Cryptosporidium parvum antigens in human fecal specimens using the Triage parasite panel enzyme immunoassay. J Clin Microbiol 38:3337–3340. Petri WA, Jr, Jackson TF, Gathiram V, Kress K, Saffer LD, Snodgrass TL, Chapman MD, Keren Z, Mirelman D. 1990. Pathogenic and nonpathogenic strains of Entamoeba histolytica can be differentiated by monoclonal antibodies to the galactose-specific adherence lectin. Infect Immun 58:1802–1806. Leo M, Haque R, Kabir M, Roy S, Lahlou RM, Mondal D, Tannich E, Petri WA, Jr. 2006. Evaluation of Entamoeba

140. Intestinal and Urogenital Parasites n 2419

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

45.

46.

47.

48.

49.

histolytica antigen and antibody point-of-care tests for the rapid diagnosis of amebiasis. J Clin Microbiol 44:4569– 4571. Korpe PS, Stott BR, Nazib F, Kabir M, Haque R, Herbein JF, Petri WA, Jr. 2012. Evaluation of a rapid point-ofcare fecal antigen detection test for Entamoeba histolytica. Am J Trop Med Hyg 86:980–981. Ong SJ, Cheng MY, Liu KH, Horng CB. 1996. Use of the ProSpecT microplate enzyme immunoassay for the detection of pathogenic and non-pathogenic Entamoeba histolytica in faecal specimens. Trans R Soc Trop Med Hyg 90:248–249. Roy S, Kabir M, Mondal D, Ali IK, Petri WA, Jr, Haque R. 2005. Real-time-PCR assay for diagnosis of Entamoeba histolytica infection. J Clin Microbiol 43:2168–2172. Stark D, van Hal S, Fotedar R, Butcher A, Marriott D, Ellis J, Harkness J. 2008. Comparison of stool antigen detection kits to PCR for diagnosis of amebiasis. J Clin Microbiol 46:1678–1681. Yau YC, Crandall I, Kain KC. 2001. Development of monoclonal antibodies which specifically recognize Entamoeba histolytica in preserved stool samples. J Clin Microbiol 39:716–719. Abd-Alla MD, Jackson TF, Reddy S, Ravdin JI. 2000. Diagnosis of invasive amebiasis by enzyme-linked immunosorbent assay of saliva to detect amebic lectin antigen and anti-lectin immunoglobulin G antibodies. J Clin Microbiol 38:2344–2347. Haque R, Mollah NU, Ali IK, Alam K, Eubanks A, Lyerly D, Petri WA, Jr. 2000. Diagnosis of amebic liver abscess and intestinal infection with the TechLab Entamoeba histolytica II antigen detection and antibody tests. J Clin Microbiol 38:3235–3239. Evangelopoulos A, Spanakos G, Patsoula E, Vakalis N, Legakis N. 2000. A nested, multiplex, PCR assay for the simultaneous detection and differentiation of Entamoeba histolytica and Entamoeba dispar in faeces. Ann Trop Med Parasitol 94:233–240. Qvarnstrom Y, James C, Xayavong M, Holloway BP, Visvesvara GS, Sriram R, da Silva AJ. 2005. Comparison of real-time PCR protocols for differential laboratory diagnosis of amebiasis. J Clin Microbiol 43:5491–5497. Rivera WL, Tachibana H, Silva-Tahat MR, Uemura H, Kanbara H. 1996. Differentiation of Entamoeba histolytica and E. dispar DNA from cysts present in stool specimens by polymerase chain reaction: its field application in the Philippines. Parasitol Res 82:585–589. Solaymani-Mohammadi S, Lam MM, Zunt JR, Petri WA, Jr. 2007. Entamoeba histolytica encephalitis diagnosed by PCR of cerebrospinal fluid. Trans R Soc Trop Med Hyg 101:311–313. Zengzhu G, Bracha R, Nuchamowitz Y, Cheng I, Mirelman D. 1999. Analysis by enzyme-linked immunosorbent assay and PCR of human liver abscess aspirates from patients in China for Entamoeba histolytica. J Clin Microbiol 37:3034–3036. Blessmann J, Buss H, Nu PA, Dinh BT, Ngo QT, Van AL, Abd Alla MD, Jackson TF, Ravdin JI, Tannich E. 2002. Real-time PCR for detection and differentiation of Entamoeba histolytica and Entamoeba dispar in fecal samples. J Clin Microbiol 40:4413–4417. Haque R, Kabir M, Noor Z, Rahman SM, Mondal D, Alam F, Rahman I, Al Mahmood A, Ahmed N, Petri WA, Jr. 2010. Diagnosis of amebic liver abscess and amebic colitis by detection of Entamoeba histolytica DNA in blood, urine, and saliva by a real-time PCR assay. J Clin Microbiol 48:2798–2801. Khairnar K, Parija SC. 2007. A novel nested multiplex polymerase chain reaction (PCR) assay for differential detection of Entamoeba histolytica, E. moshkovskii and E. dispar

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2419

50.

51.

52.

53.

54.

55.

56.

57.

58.

59.

60.

61.

62. 63.

64. 65. 66.

DNA in stool samples. BMC Microbiol 7:47. doi:10.1186/ 1471-2180-7-47. Santos HL, Bandyopadhyay K, Bandea R, Peralta RH, Peralta JM, Da Silva AJ. 2013. LUMINEX®: a new technology for the simultaneous identification of five Entamoeba spp. commonly found in human stools. Parasit Vectors 6:69. doi:10.1186/1756-3305-6-69. Verweij JJ, Blange RA, Templeton K, Schinkel J, Brienen EA, van Rooyen MA, van Lieshout L, Polderman AM. 2004. Simultaneous detection of Entamoeba histolytica, Giardia lamblia, and Cryptosporidium parvum in fecal samples by using multiplex real-time PCR. J Clin Microbiol 42:1220–1223. Stark D, Al-Qassab SE, Barratt JL, Stanley K, Roberts T, Marriott D, Harkness J, Ellis JT. 2011. Evaluation of multiplex tandem real-time PCR for detection of Cryptosporidium spp., Dientamoeba fragilis, Entamoeba histolytica, and Giardia intestinalis in clinical stool samples. J Clin Microbiol 49:257–262. Claas E, Burnham CA, Mazulli T, Templeton K, Topin F. 2013. Performance of the xTAG® Gastrointestinal Pathogen Panel (GPP), a multiplex molecular assay for simultaneous detection of bacterial, viral, and parasitic causes of infectious gastroenteritis. J Microbiol Biotechnol 23:1041–1045. Haque R, Ali IK, Akther S, Petri WA, Jr. 1998. Comparison of PCR, isoenzyme analysis, and antigen detection for diagnosis of Entamoeba histolytica infection. J Clin Microbiol 36:449–452. Paglia MG, Visca P. 2004. An improved PCR-based method for detection and differentiation of Entamoeba histolytica and Entamoeba dispar in formalin-fixed stools. Acta Trop 92:273–277. Troll H, Marti H, Weiss N. 1997. Simple differential detection of Entamoeba histolytica and Entamoeba dispar in fresh stool specimens by sodium acetate-acetic acidformalin concentration and PCR. J Clin Microbiol 35:1701– 1705. Verweij JJ, Blotkamp J, Brienen EA, Aguirre A, Polderman AM. 2000. Differentiation of Entamoeba histolytica and Entamoeba dispar cysts using polymerase chain reaction on DNA isolated from faeces with spin columns. Eur J Clin Microbiol Infect Dis 19:358–361. Monteiro L, Bonnemaison D, Vekris A, Petry KG, Bonnet J, Vidal R, Cabrita J, Megraud F. 1997. Complex polysaccharides as PCR inhibitors in feces: Helicobacter pylori model. J Clin Microbiol 35:995–998. Joyce MP, Ravdin JI. 1988. Antigens of Entamoeba histolytica recognized by immune sera from liver abscess patients. Am J Trop Med Hyg 38:74–80. Haque R, Ikm A, Petri WA, Jr. 2000. Salivary antilectin IgA antibodies in a cohort of children residing in an endemic area of Bangladesh. Arch Med Res 31:S41–S43. Stanley SL, Jr. 2001. Protective immunity to amebiasis: new insights and new challenges. J Infect Dis 184:504– 506. The Medical Letter. 2013. Drugs for Parasitic Infections, 3rd ed. The Medical Letter, Inc, New Rochelle, NY. Upcroft P, Upcroft JA. 2001. Drug targets and mechanisms of resistance in the anaerobic protozoa. Clin Microbiol Rev 14:150–164. Cavalier-Smith T. 1998. A revised six-kingdom system of life. Biol Rev Camb Philos Soc 73:203–266. Silberman JD, Sogin ML, Leipe DD, Clark CG. 1996. Human parasite finds taxonomic home. Nature 380:398. Arisue N, Hashimoto T, Yoshikawa H, Nakamura Y, Nakamura G, Nakamura F, Yano TA, Hasegawa M. 2002. Phylogenetic position of Blastocystis hominis and of stramenopiles inferred from multiple molecular sequence data. J Eukaryot Microbiol 49:42–53.

2420 n

PARASITOLOGY

67. Hoevers JD, Snowden KF. 2005. Analysis of the ITS region and partial ssu and lsu rRNA genes of Blastocystis and Proteromonas lacertae. Parasitology 131:187–196. 68. Tan KS. 2008. New insights on classification, identification, and clinical relevance of Blastocystis spp. Clin Microbiol Rev 21:639–665. 69. Clark CG. 1997. Extensive genetic diversity in Blastocystis hominis. Mol Biochem Parasitol 87:79–83. 70. Mansour NS, Mikhail EM, el Masry NA, Sabry AG, Mohareb EW. 1995. Biochemical characterisation of human isolates of Blastocystis hominis. J Med Microbiol 42:304–307. 71. Noel C, Dufernez F, Gerbod D, Edgcomb VP, DelgadoViscogliosi P, Ho LC, Singh M, Wintjens R, Sogin ML, Capron M, Pierce R, Zenner L, Viscogliosi E. 2005. Molecular phylogenies of Blastocystis isolates from different hosts: implications for genetic diversity, identification of species, and zoonosis. J Clin Microbiol 43:348–355. 72. Leelayoova S, Rangsin R, Taamasri P, Naaglor T, Thathaisong U, Mungthin M. 2004. Evidence of waterborne transmission of Blastocystis hominis. Am J Trop Med Hyg 70:658–662. 73. Yoshikawa H, Abe N, Iwasawa M, Kitano S, Nagano I, Wu Z, Takahashi Y. 2000. Genomic analysis of Blastocystis hominis strains isolated from two long-term health care facilities. J Clin Microbiol 38:1324–1330. 74. Moe KT, Singh M, Howe J, Ho LC, Tan SW, Ng GC, Chen XQ, Yap EH. 1996. Observations on the ultrastructure and viability of the cystic stage of Blastocystis hominis from human feces. Parasitol Res 82:439–444. 75. Markell EK, Udkow MP. 1986. Blastocystis hominis: pathogen or fellow traveler? Am J Trop Med Hyg 35:1023–1026. 76. Sheehan DJ, Raucher BG, McKitrick JC. 1986. Association of Blastocystis hominis with signs and symptoms of human disease. J Clin Microbiol 24:548–550. 77. Zierdt CH. 1991. Blastocystis hominis—past and future. Clin Microbiol Rev 4:61–79. 78. Horiki N, Kaneda Y, Maruyama M, Fujita Y, Tachibana H. 1999. Intestinal blockage by carcinoma and Blastocystis hominis infection. Am J Trop Med Hyg 60:400–402. 79. Udkow MP, Markell EK. 1993. Blastocystis hominis: prevalence in asymptomatic versus symptomatic hosts. J Infect Dis 168:242–244. 80. Albrecht H, Stellbrink HJ, Koperski K, Greten H. 1995. Blastocystis hominis in human immunodeficiency virusrelated diarrhea. Scand J Gastroenterol 30:909–914. 81. Rapeeporn Y, Warunee N, Nuttapong W, Sompong S, Rachada K. 2006. Infection of Blastocystis hominis in primary schoolchildren from Nakhon Pathom province, Thailand. Trop Biomed 23:117–122. 82. Leelayoova S, Taamasri P, Rangsin R, Naaglor T, Thathaisong U, Mungthin M. 2002. In-vitro cultivation: a sensitive method for detecting Blastocystis hominis. Ann Trop Med Parasitol 96:803–807. 83. Garavelli PL, Zierdt CH, Fleisher TA, Liss H, Nagy B. 1995. Serum antibody detected by fluorescent antibody test in patients with symptomatic Blastocystis hominis infection. Recenti Prog Med 86:398–400. 84. Zierdt CH, Zierdt WS, Nagy B. 1995. Enzyme-linked immunosorbent assay for detection of serum antibody to Blastocystis hominis in symptomatic infections. J Parasitol 81:127–129. 85. Kaneda Y, Horiki N, Cheng X, Tachibana H, Tsutsumi Y. 2000. Serologic response to Blastocystis hominis infection in asymptomatic individuals. Tokai J Exp Clin Med 25:51– 56. 86. Stensvold CR, Arendrup MC, Jespersgaard C, Molbak K, Nielsen HV. 2007. Detecting Blastocystis using parasitologic and DNA-based methods: a comparative study. Diagn Microbiol Infect Dis 59:303–307.

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2420

87. Stensvold CR, Traub RJ, von Samson-Himmelstjerna G, Jespersgaard C, Nielsen HV, Thompson RC. 2007. Blastocystis: subtyping isolates using pyrosequencing technology. Exp Parasitol 116:111–119. 88. Stensvold R, Brillowska-Dabrowska A, Nielsen HV, Arendrup MC. 2006. Detection of Blastocystis hominis in unpreserved stool specimens by using polymerase chain reaction. J Parasitol 92:1081–1087. 89. Haresh K, Suresh K, Khairul AA, Saminathan S. 1999. Isolate resistance of Blastocystis hominis to metronidazole. Trop Med Int Health 4:274–277. 90. Feng Y, Xiao L. 2011. Zoonotic potential and molecular epidemiology of Giardia species and giardiasis. Clin Microbiol Rev 24:110–140. 91. Guy RA, Xiao C, Horgen PA. 2004. Real-time PCR assay for detection and genotype differentiation of Giardia lamblia in stool specimens. J Clin Microbiol 42:3317–3320. 92. Rendtorff RC. 1978. The experimental transmission of Giardia lamblia among volunteer subjects, p 64–81. In Jacubowski W, Hoff JC (ed), Waterborne Transmission of Giardiasis 1978. EPA/600/9-79-001. U.S. Environmental Protection Agency, Washington, DC. 93. Panosian CB. 1988. Parasitic diarrhea. Infect Dis Clin North Am 2:685–703. 94. Olson ME, Ceri H, Morck DW. 2000. Giardia vaccination. Parasitol Today 16:213–217. 95. Monis PT, Andrews RH, Mayrhofer G, Ey PL. 1999. Molecular systematics of the parasitic protozoan Giardia intestinalis. Mol Biol Evol 16:1135–1144. 96. Paintlia AS, Descoteaux S, Spencer B, Chakraborti A, Ganguly NK, Mahajan RC, Samuelson J. 1998. Giardia lamblia groups A and B among young adults in India. Clin Infect Dis 26:190–191. 97. Paintlia AS, Paintlia MK, Mahajan RC, Chakraborti A, Ganguly NK. 1999. A DNA-based probe for differentiation of Giardia lamblia group A and B isolates from northern India. Clin Infect Dis 28:1178–1180. 98. Carroccio A, Montalto G, Iacono G, Ippolito S, Soresi M, Notarbartolo A. 1997. Secondary impairment of pancreatic function as a cause of severe malabsorption in intestinal giardiasis: a case report. Am J Trop Med Hyg 56:599–602. 99. Garcia LS, Shimizu RY. 1997. Evaluation of nine immunoassay kits (enzyme immunoassay and direct fluorescence) for detection of Giardia lamblia and Cryptosporidium parvum in human fecal specimens. J Clin Microbiol 35:1526–1529. 100. Johnston SP, Ballard MM, Beach MJ, Causer L, Wilkins PP. 2003. Evaluation of three commercial assays for detection of Giardia and Cryptosporidium organisms in fecal specimens. J Clin Microbiol 41:623–626. 101. Katanik MT, Schneider SK, Rosenblatt JE, Hall GS, Procop GW. 2001. Evaluation of ColorPAC Giardia/ Cryptosporidium rapid assay and ProSpecT Giardia/Cryptosporidium microplate assay for detection of Giardia and Cryptosporidium in fecal specimens. J Clin Microbiol 39:4523–4525. 102. Maraha B, Buiting AG. 2000. Evaluation of four enzyme immunoassays for the detection of Giardia lamblia antigen in stool specimens. Eur J Clin Microbiol Infect Dis 19:485– 487. 103. Youn S, Kabir M, Haque R, Petri WA, Jr. 2009. Evaluation of a screening test for detection of Giardia and Cryptosporidium parasites. J Clin Microbiol 47:451–452. 104. Goni P, Martin B, Villacampa M, Garcia A, Seral C, Castillo FJ, Clavel A. 2012. Evaluation of an immunochromatographic dip strip test for simultaneous detection of Cryptosporidium spp, Giardia duodenalis, and Entamoeba histolytica antigens in human faecal samples. Eur J Clin Microbiol Infect Dis 31:2077–2082.

140. Intestinal and Urogenital Parasites n 2421 105. Garcia LS, Shimizu RY. 2000. Detection of Giardia lamblia and Cryptosporidium parvum antigens in human fecal specimens using the ColorPAC combination rapid solidphase qualitative immunochromatographic assay. J Clin Microbiol 38:1267–1268. 106. Christy NC, Hencke JD, Escueta-De Cadiz A, Nazib F, von Thien H, Yagita K, Ligaba S, Haque R, Nozaki T, Tannich E, Herbein JF, Petri WA, Jr. 2012. Multisite performance evaluation of an enzyme-linked immunosorbent assay for detection of Giardia, Cryptosporidium, and Entamoeba histolytica antigens in human stool. J Clin Microbiol 50:1762–1763. 107. Garcia LS, Garcia JP. 2006. Detection of Giardia lamblia antigens in human fecal specimens by a solid-phase qualitative immunochromatographic assay. J Clin Microbiol 44:4587–4588. 108. Hanson KL, Cartwright CP. 2001. Use of an enzyme immunoassay does not eliminate the need to analyze multiple stool specimens for sensitive detection of Giardia lamblia. J Clin Microbiol 39:474–477. 109. Garcia LS, Shimizu RY, Novak S, Carroll M, Chan F. 2003. Commercial assay for detection of Giardia lamblia and Cryptosporidium parvum antigens in human fecal specimens by rapid solid-phase qualitative immunochromatography. J Clin Microbiol 41:209–212. 110. Pillai DR, Kain KC. 1999. Immunochromatographic strip-based detection of Entamoeba histolytica-E. dispar and Giardia lamblia coproantigen. J Clin Microbiol 37:3017– 3019. 111. Sharp SE, Suarez CA, Duran Y, Poppiti RJ. 2001. Evaluation of the Triage Micro Parasite Panel for detection of Giardia lamblia, Entamoeba histolytica/Entamoeba dispar, and Cryptosporidium parvum in patient stool specimens. J Clin Microbiol 39:332–334. 112. Minak J, Kabir M, Mahmud I, Liu Y, Liu L, Haque R, Petri WA, Jr. 2012. Evaluation of rapid antigen pointof-care tests for detection of Giardia and Cryptosporidium species in human fecal specimens. J Clin Microbiol 50:154– 156. 113. Amar CF, Dear PH, Pedraza-Diaz S, Looker N, Linnane E, McLauchlin J. 2002. Sensitive PCR-restriction fragment length polymorphism assay for detection and genotyping of Giardia duodenalis in human feces. J Clin Microbiol 40:446–452. 114. Nantavisai K, Mungthin M, Tan-ariya P, Rangsin R, Naaglor T, Leelayoova S. 2007. Evaluation of the sensitivities of DNA extraction and PCR methods for detection of Giardia duodenalis in stool specimens. J Clin Microbiol 45:581–583. 115. Wang Z, Vora GJ, Stenger DA. 2004. Detection and genotyping of Entamoeba histolytica, Entamoeba dispar, Giardia lamblia, and Cryptosporidium parvum by oligonucleotide microarray. J Clin Microbiol 42:3262–3271. 116. Ng CT, Gilchrist CA, Lane A, Roy S, Haque R, Houpt ER. 2005. Multiplex real-time PCR assay using Scorpion probes and DNA capture for genotype-specific detection of Giardia lamblia on fecal samples. J Clin Microbiol 43:1256–1260. 117. Rochelle PA, De Leon R, Stewart MH, Wolfe RL. 1997. Comparison of primers and optimization of PCR conditions for detection of Cryptosporidium parvum and Giardia lamblia in water. Appl Environ Microbiol 63:106–114. 118. Weiss JB. 1995. DNA probes and PCR for diagnosis of parasitic infections. Clin Microbiol Rev 8:113–130. 119. Gardner TB, Hill DR. 2001. Treatment of giardiasis. Clin Microbiol Rev 14:114–128. 120. Marshall MM, Naumovitz D, Ortega Y, Sterling CR. 1997. Waterborne protozoan pathogens. Clin Microbiol Rev 10:67–85. 121. Sousa MC, Poiares-Da-Silva J. 1999. A new method for assessing metronidazole susceptibility of Giardia lamblia

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2421

122.

123.

124.

125.

126.

127.

128.

129. 130.

131. 132.

133. 134.

135.

136.

137.

138.

139.

140.

141.

trophozoites. Antimicrob Agents Chemother 43:2939– 2942. Gerbod D, Edgcomb VP, Noel C, Zenner L, Wintjens R, Delgado-Viscogliosi P, Holder ME, Sogin ML, Viscogliosi E. 2001. Phylogenetic position of the trichomonad parasite of turkeys, Histomonas meleagridis (Smith) Tyzzer, inferred from small subunit rRNA sequence. J Eukaryot Microbiol 48:498–504. Silberman JD, Clark CG, Sogin ML. 1996. Dientamoeba fragilis shares a recent common evolutionary history with the trichomonads. Mol Biochem Parasitol 76:311–314. Burrows RB, Swerdlow MA. 1956. Enterobius vermicularis as a probable vector of Dientamoeba fragilis. Am J Trop Med Hyg 5:258–265. Johnson EH, Windsor JJ, Clark CG. 2004. Emerging from obscurity: biological, clinical, and diagnostic aspects of Dientamoeba fragilis. Clin Microbiol Rev 17:553–570. Munasinghe VS, Vella NG, Ellis JT, Windsor PA, Stark D. 2013. Cyst formation and faecal-oral transmission of Dientamoeba fragilis—the missing link in the life cycle of an emerging pathogen. Int J Parasitol 43:879–883. Clark CG, Roser D, Stensvold CR. 2014. Transmission of Dientamoeba fragilis: pinworm or cysts? Trends Parasitol 30:136–140. Yang J, Scholten T. 1977. Dientamoeba fragilis: a review with notes on its epidemiology, pathogenicity, mode of transmission, and diagnosis. Am J Trop Med Hyg 26:16– 22. Johnson JA, Clark CG. 2000. Cryptic genetic diversity in Dientamoeba fragilis. J Clin Microbiol 38:4653–4654. Peek R, Reedeker FR, van Gool T. 2004. Direct amplification and genotyping of Dientamoeba fragilis from human stool specimens. J Clin Microbiol 42:631–635. Butler WP. 1996. Dientamoeba fragilis. An unusual intestinal pathogen. Dig Dis Sci 41:1811–1813. Schure JM, de Vries M, Weel JF, van Roon EN, Faber TE. 2013. Symptoms and treatment of Dientamoeba fragilis infection in children, a retrospective study. Pediatr Infect Dis J 32:e148–e150. Talis B, Stein B, Lengy J. 1971. Dientamoeba fragilis in human feces and bile. Isr J Med Sci 7:1063–1069. Cuffari C, Oligny L, Seidman EG. 1998. Dientamoeba fragilis masquerading as allergic colitis. J Pediatr Gastroenterol Nutr 26:16–20. Lainson R, da Silva BA. 1999. Intestinal parasites of some diarrhoeic HIV-seropositive individuals in North Brazil, with particular reference to Isospora belli Wenyon, 1923 and Dientamoeba fragilis Jepps & Dobell, 1918. Mem Inst Oswaldo Cruz 94:611–613. Schwartz MD, Nelson ME. 2003. Dientamoeba fragilis infection presenting to the emergency department as acute appendicitis. J Emerg Med 25:17–21. Gray TJ, Kwan YL, Phan T, Robertson G, Cheong EY, Gottlieb T. 2013. Dientamoeba fragilis: a family cluster of disease associated with marked peripheral eosinophilia. Clin Infect Dis 57:845–848. Chan F, Stewart N, Guan M, Robb I, Fuite L, Chan I, Diaz-Mitoma F, King J, MacDonald N, Mackenzie A. 1996. Prevalence of Dientamoeba fragilis antibodies in children and recognition of a 39 kDa immunodominant protein antigen of the organism. Eur J Clin Microbiol Infect Dis 15:950–954. Chan FT, Guan MX, Mackenzie AM. 1993. Application of indirect immunofluorescence to detection of Dientamoeba fragilis trophozoites in fecal specimens. J Clin Microbiol 31:1710–1714. Stark D, Beebe N, Marriott D, Ellis J, Harkness J. 2005. Detection of Dientamoeba fragilis in fresh stool specimens using PCR. Int J Parasitol 35:57–62. Stark D, Beebe N, Marriott D, Ellis J, Harkness J. 2006. Evaluation of three diagnostic methods, including real-

2422 n

142.

143.

144. 145.

146.

147.

148.

149.

150.

151.

152.

153. 154.

155.

156.

PARASITOLOGY

time PCR, for detection of Dientamoeba fragilis in stool specimens. J Clin Microbiol 44:232–235. Calderaro A, Gorrini C, Montecchini S, Peruzzi S, Piccolo G, Rossi S, Gargiulo F, Manca N, Dettori G, Chezzi C. 2010. Evaluation of a real-time polymerase chain reaction assay for the laboratory diagnosis of giardiasis. Diagn Microbiol Infect Dis 66:261–267. Girginkardesler N, Coskun S, Cuneyt Balcioglu I, Ertan P, Ok UZ. 2003. Dientamoeba fragilis, a neglected cause of diarrhea, successfully treated with secnidazole. Clin Microbiol Infect 9:110–113. Poole DN, McClelland RS. 2013. Global epidemiology of Trichomonas vaginalis. Sex Transm Infect 89:418–422. Schwebke JR, Hook EW, III. 2003. High rates of Trichomonas vaginalis among men attending a sexually transmitted diseases clinic: implications for screening and urethritis management. J Infect Dis 188:465–468. Munson KL, Napierala M, Munson E, Schell RF, Kramme T, Miller C, Hryciuk JE. 2013. Screening of male patients for Trichomonas vaginalis with transcriptionmediated amplification in a community with a high prevalence of sexually transmitted infection. J Clin Microbiol 51:101–104. Andrea SB, Chapin KC. 2011. Comparison of Aptima Trichomonas vaginalis transcription-mediated amplification assay and BD Affirm VPIII for detection of T. vaginalis in symptomatic women: performance parameters and epidemiological implications. J Clin Microbiol 49:866– 869. Ginocchio CC, Chapin K, Smith JS, Aslanzadeh J, Snook J, Hill CS, Gaydos CA. 2012. Prevalence of Trichomonas vaginalis and coinfection with Chlamydia trachomatis and Neisseria gonorrhoeae in the United States as determined by the Aptima Trichomonas vaginalis nucleic acid amplification assay. J Clin Microbiol 50:2601–2608. Stark JR, Judson G, Alderete JF, Mundodi V, Kucknoor AS, Giovannucci EL, Platz EA, Sutcliffe S, Fall K, Kurth T, Ma J, Stampfer MJ, Mucci LA. 2009. Prospective study of Trichomonas vaginalis infection and prostate cancer incidence and mortality: Physicians’ Health Study. J Natl Cancer Inst 101:1406–1411. Sutcliffe S, Giovannucci E, Alderete JF, Chang TH, Gaydos CA, Zenilman JM, De Marzo AM, Willett WC, Platz EA. 2006. Plasma antibodies against Trichomonas vaginalis and subsequent risk of prostate cancer. Cancer Epidemiol Biomarkers Prev 15:939–945. Mitteregger D, Aberle SW, Makristathis A, Walochnik J, Brozek W, Marberger M, Kramer G. 2012. High detection rate of Trichomonas vaginalis in benign hyperplastic prostatic tissue. Med Microbiol Immunol 201:113–116. Neace CJ, Alderete JF. 2013. Epitopes of the highly immunogenic Trichomonas vaginalis α-actinin are serodiagnostic targets for both women and men. J Clin Microbiol 51:2483–2490. Schwebke JR, Burgess D. 2004. Trichomoniasis. Clin Microbiol Rev 17:794–803. Cotch MF, Pastorek JG, II, Nugent RP, Yerg DE, Martin DH, Eschenbach DA, The Vaginal Infections and Prematurity Study Group. 1991. Demographic and behavioral predictors of Trichomonas vaginalis infection among pregnant women. Obstet Gynecol 78:1087–1092. Minkoff H, Grunebaum AN, Schwarz RH, Feldman J, Cummings M, Crombleholme W, Clark L, Pringle G, McCormack WM. 1984. Risk factors for prematurity and premature rupture of membranes: a prospective study of the vaginal flora in pregnancy. Am J Obstet Gynecol 150:965–972. Soper DE, Bump RC, Hurt WG. 1990. Bacterial vaginosis and trichomoniasis vaginitis are risk factors for cuff cellulitis after abdominal hysterectomy. Am J Obstet Gynecol 163:1016–1021.

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2422

157. Jackson DJ, Rakwar JP, Bwayo JJ, Kreiss JK, Moses S. 1997. Urethral Trichomonas vaginalis infection and HIV-1 transmission. Lancet 350:1076. 158. Laga M, Manoka A, Kivuvu M, Malele B, Tuliza M, Nzila N, Goeman J, Behets F, Batter V, Alary M, et al. 1993. Non-ulcerative sexually transmitted diseases as risk factors for HIV-1 transmission in women: results from a cohort study. AIDS 7:95–102. 159. Hobbs MM, Kazembe P, Reed AW, Miller WC, Nkata E, Zimba D, Daly CC, Chakraborty H, Cohen MS, Hoffman I. 1999. Trichomonas vaginalis as a cause of urethritis in Malawian men. Sex Transm Dis 26:381–387. 160. McLaren LC, Davis LE, Healy GR, James CG. 1983. Isolation of Trichomonas vaginalis from the respiratory tract of infants with respiratory disease. Pediatrics 71:888–890. 161. Cornelius DC, Robinson DA, Muzny CA, Mena LA, Aanensen DM, Lushbaugh WB, Meade JC. 2012. Genetic characterization of Trichomonas vaginalis isolates by use of multilocus sequence typing. J Clin Microbiol 50:3293–3300. 162. Krieger JN, Tam MR, Stevens CE, Nielsen IO, Hale J, Kiviat NB, Holmes KK. 1988. Diagnosis of trichomoniasis. Comparison of conventional wet-mount examination with cytologic studies, cultures, and monoclonal antibody staining of direct specimens. JAMA 259:1223–1227. 163. Heine RP, Wiesenfeld HC, Sweet RL, Witkin SS. 1997. Polymerase chain reaction analysis of distal vaginal specimens: a less invasive strategy for detection of Trichomonas vaginalis. Clin Infect Dis 24:985–987. 164. Schwebke JR, Morgan SC, Pinson GB. 1997. Validity of self-obtained vaginal specimens for diagnosis of trichomoniasis. J Clin Microbiol 35:1618–1619. 165. Hook EW, III. 1999. Trichomonas vaginalis—no longer a minor STD. Sex Transm Dis 26:388–389. 166. Borchardt KA, Smith RF. 1991. An evaluation of an InPouch™ TV culture method for diagnosing Trichomonas vaginalis infection. Genitourin Med 67:149–152. 167. Draper D, Parker R, Patterson E, Jones W, Beutz M, French J, Borchardt K, McGregor J. 1993. Detection of Trichomonas vaginalis in pregnant women with the InPouch TV culture system. J Clin Microbiol 31:1016–1018. 168. Schwebke JR, Venglarik MF, Morgan SC. 1999. Delayed versus immediate bedside inoculation of culture media for diagnosis of vaginal trichomonosis. J Clin Microbiol 37:2369–2370. 169. Adu-Sarkodie Y, Opoku BK, Danso KA, Weiss HA, Mabey D. 2004. Comparison of latex agglutination, wet preparation, and culture for the detection of Trichomonas vaginalis. Sex Transm Infect 80:201–203. 170. Campbell L, Woods V, Lloyd T, Elsayed S, Church DL. 2008. Evaluation of the OSOM Trichomonas rapid test versus wet preparation examination for detection of Trichomonas vaginalis vaginitis in specimens from women with a low prevalence of infection. J Clin Microbiol 46:3467–3469. 171. Huppert JS, Batteiger BE, Braslins P, Feldman JA, Hobbs MM, Sankey HZ, Sena AC, Wendel KA. 2005. Use of an immunochromatographic assay for rapid detection of Trichomonas vaginalis in vaginal specimens. J Clin Microbiol 43:684–687. 172. Huppert JS, Mortensen JE, Reed JL, Kahn JA, Rich KD, Miller WC, Hobbs MM. 2007. Rapid antigen testing compares favorably with transcription-mediated amplification assay for the detection of Trichomonas vaginalis in young women. Clin Infect Dis 45:194–198. 173. Pattullo L, Griffeth S, Ding L, Mortensen J, Reed J, Kahn J, Huppert J. 2009. Stepwise diagnosis of Trichomonas vaginalis infection in adolescent women. J Clin Microbiol 47:59–63. 174. Kurth A, Whittington WL, Golden MR, Thomas KK, Holmes KK, Schwebke JR. 2004. Performance of a new,

140. Intestinal and Urogenital Parasites n 2423

175.

176.

177.

178.

179.

180.

181.

182.

183.

184.

185.

186.

187.

188.

189.

rapid assay for detection of Trichomonas vaginalis. J Clin Microbiol 42:2940–2943. Pillay A, Lewis J, Ballard RC. 2004. Evaluation of Xenostrip-Tv, a rapid diagnostic test for Trichomonas vaginalis infection. J Clin Microbiol 42:3853–3856. Jones HE, Lippman SA, Caiaffa-Filho HH, Young T, van de Wijgert JH. 2013. Performance of a rapid selftest for detection of Trichomonas vaginalis in South Africa and Brazil. J Clin Microbiol 51:1037–1039. Pearce DM, Styles DN, Hardick JP, Gaydos CA. 2013. A new rapid molecular point-of-care assay for Trichomonas vaginalis: preliminary performance data. Sex Transm Infect 89:495–497. Brown HL, Fuller DD, Jasper LT, Davis TE, Wright JD. 2004. Clinical evaluation of Affirm VPIII in the detection and identification of Trichomonas vaginalis, Gardnerella vaginalis, and Candida species in vaginitis/vaginosis. Infect Dis Obstet Gynecol 12:17–21. Hardick J, Yang S, Lin S, Duncan D, Gaydos C. 2003. Use of the Roche LightCycler instrument in a real-time PCR for Trichomonas vaginalis in urine samples from females and males. J Clin Microbiol 41:5619–5622. Lin PR, Shaio MF, Liu JY. 1997. One-tube, nested-PCR assay for the detection of Trichomonas vaginalis in vaginal discharges. Ann Trop Med Parasitol 91:61–65. Munson E, Napierala M, Olson R, Endes T, Block T, Hryciuk JE, Schell RF. 2008. Impact of Trichomonas vaginalis transcription-mediated amplification-based analyte-specific-reagent testing in a metropolitan setting of high sexually transmitted disease prevalence. J Clin Microbiol 46:3368–3374. Nye MB, Schwebke JR, Body BA. 2009. Comparison of APTIMA Trichomonas vaginalis transcription-mediated amplification to wet mount microscopy, culture, and polymerase chain reaction for diagnosis of trichomoniasis in men and women. Am J Obstet Gynecol 200:188.e1– 188.e7. doi:10.1016/j.ajog.2008.10.005. Riley DE, Roberts MC, Takayama T, Krieger JN. 1992. Development of a polymerase chain reaction-based diagnosis of Trichomonas vaginalis. J Clin Microbiol 30:465– 472. Kaydos SC, Swygard H, Wise SL, Sena AC, Leone PA, Miller WC, Cohen MS, Hobbs MM. 2002. Development and validation of a PCR-based enzyme-linked immunosorbent assay with urine for use in clinical research settings to detect Trichomonas vaginalis in women. J Clin Microbiol 40:89–95. Lawing LF, Hedges SR, Schwebke JR. 2000. Detection of trichomonosis in vaginal and urine specimens from women by culture and PCR. J Clin Microbiol 38:3585– 3588. Kaydos-Daniels SC, Miller WC, Hoffman I, Banda T, Dzinyemba W, Martinson F, Cohen MS, Hobbs MM. 2003. Validation of a urine-based PCR–enzyme-linked immunosorbent assay for use in clinical research settings to detect Trichomonas vaginalis in men. J Clin Microbiol 41:318–323. Schwebke JR, Lawing LF. 2002. Improved detection by DNA amplification of Trichomonas vaginalis in males. J Clin Microbiol 40:3681–3683. Hardick A, Hardick J, Wood BJ, Gaydos C. 2006. Comparison between the Gen-Probe transcription-mediated amplification Trichomonas vaginalis research assay and real-time PCR for Trichomonas vaginalis detection using a Roche LightCycler instrument with female self-obtained vaginal swab samples and male urine samples. J Clin Microbiol 44:4197–4199. Schwebke JR, Hobbs MM, Taylor SN, Sena AC, Catania MG, Weinbaum BS, Johnson AD, Getman DK, Gaydos CA. 2011. Molecular testing for Trichomonas vagi-

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2423

190.

191.

192.

193.

194.

195.

196.

197.

198.

199.

200. 201.

202.

203.

204.

205.

nalis in women: results from a prospective U.S. clinical trial. J Clin Microbiol 49:4106–4111. Bandea CI, Joseph K, Secor EW, Jones LA, Igietseme JU, Sautter RL, Hammerschlag MR, Fajman NN, Girardet RG, Black CM. 2013. Development of PCR assays for detection of Trichomonas vaginalis in urine specimens. J Clin Microbiol 51:1298–1300. Cosentino LA, Campbell T, Jett A, Macio I, Zamborsky T, Cranston RD, Hillier SL. 2012. Use of nucleic acid amplification testing for diagnosis of anorectal sexually transmitted infections. J Clin Microbiol 50:2005–2008. Lee SJ, Park DC, Lee DS, Choe HS, Cho YH. 2012. Evaluation of Seeplex® STD6 ACE Detection kit for the diagnosis of six bacterial sexually transmitted infections. J Infect Chemother 18:494–500. Workowski KA, Berman S, Centers for Disease Control and Prevention (CDC). 2010. Sexually transmitted diseases treatment guidelines, 2010. MMWR Recomm Rep 59:1–110. Cudmore SL, Delgaty KL, Hayward-McClelland SF, Petrin DP, Garber GE. 2004. Treatment of infections caused by metronidazole-resistant Trichomonas vaginalis. Clin Microbiol Rev 17:783–793. Klebanoff MA, Carey JC, Hauth JC, Hillier SL, Nugent RP, Thom EA, Ernest JM, Heine RP, Wapner RJ, Trout W, Moawad A, Leveno KJ, Miodovnik M, Sibai BM, Van Dorsten JP, Dombrowski MP, O’Sullivan MJ, Varner M, Langer O, McNellis D, Roberts JM. 2001. Failure of metronidazole to prevent preterm delivery among pregnant women with asymptomatic Trichomonas vaginalis infection. N Engl J Med 345:487–493. Sobel JD, Nagappan V, Nyirjesy P. 1999. Metronidazoleresistant vaginal trichomoniasis—an emerging problem. N Engl J Med 341:292–293. Borchardt KA, Li Z, Zhang MZ, Shing H. 1996. An in vitro metronidazole susceptibility test for trichomoniasis using the InPouch TV™ test. Genitourin Med 72:132– 135. Meri T, Jokiranta TS, Suhonen L, Meri S. 2000. Resistance of Trichomonas vaginalis to metronidazole: report of the first three cases from Finland and optimization of in vitro susceptibility testing under various oxygen concentrations. J Clin Microbiol 38:763–767. Kawamura O, Kon Y, Naganuma A, Iwami T, Maruyama H, Yamada T, Sonobe K, Horikoshi T, Kusano M, Mori M. 2001. Retortamonas intestinalis in the pancreatic juice of a patient with small nodular lesions of the main pancreatic duct. Gastrointest Endosc 53:508–510. Schuster FL, Ramirez-Avila L. 2008. Current world status of Balantidium coli. Clin Microbiol Rev 21:626–638. Struder-Kypke MC, Kornilova OA, Lynn DH. 2007. Phylogeny of trichostome ciliates (Ciliophora, Litostomatea) endosymbiotic in the Yakut horse (Equus caballus). Eur J Protistol 43:319–328. Struder-Kypke MC, Wright AD, Foissner W, Chatzinotas A, Lynn DH. 2006. Molecular phylogeny of litostome ciliates (Ciliophora, Litostomatea) with emphasis on freeliving haptorian genera. Protist 157:261–278. Esteban JG, Aguirre C, Angles R, Ash LR, Mas-Coma S. 1998. Balantidiasis in Aymara children from the northern Bolivian Altiplano. Am J Trop Med Hyg 59:922–927. Solaymani-Mohammadi S, Rezaian M, Hooshyar H, Mowlavi GR, Babaei Z, Anwar MA. 2004. Intestinal protozoa in wild boars (Sus scrofa) in western Iran. J Wildl Dis 40:801–803. Yazar S, Altuntas F, Sahin I, Atambay M. 2004. Dysentery caused by Balantidium coli in a patient with nonHodgkin’s lymphoma from Turkey. World J Gastroenterol 10:458–459.

2424 n

PARASITOLOGY

206. Castro J, Vazquez-Iglesias JL, Arnal-Monreal F. 1983. Dysentery caused by Balantidium coli—report of two cases. Endoscopy 15:272–274. 207. Currie AR. 1990. Human balantidiasis. A case report. S Afr J Surg 28:23–25. 208. Knight R. 1978. Giardiasis, isosporiasis and balantidiasis. Clin Gastroenterol 7:31–47. 209. Ladas SD, Savva S, Frydas A, Kaloviduris A, Hatzioannou J, Raptis S. 1989. Invasive balantidiasis presented

MCM 11th Edition X : MCM11$C140 03-22-15 03:39:42

Page 2424

as chronic colitis and lung involvement. Dig Dis Sci 34:1621–1623. 210. Dhawan S, Jain D, Mehta VS. 2013. Balantidium coli: an unrecognized cause of vertebral osteomyelitis and myelopathy. J Neurosurg Spine 18:310–313. 211. Abdolrasouli A, Croucher A, Roushan A, Gaydos C. 2013. Bilateral conjunctivitis due to Trichomonas vaginalis without genital infection: an unusual presentation. J Clin Microbiol 51:3157–3159.

Cystoisospora, Cyclospora, and Sarcocystis* DAVID S. LINDSAY AND LOUIS M. WEISS

141 Cystoisospora, Cyclospora, and Sarcocystis are intestinal coccidia of humans (Fig. 1 to 5). They have varied life cycles, epidemiologies, treatment requirements, and diagnostic methods. Oocysts of these coccidia are found in the feces of humans (Table 1), and diagnosis is based ultimately on demonstrating oocysts (Cystoisospora or Cyclospora) or sporocysts (Sarcocystis) in human stool samples.

TAXONOMY Cystoisospora, Cyclospora, and Sarcocystis are in the phylum Apicomplexa, class Coccidea, order Eimeriida. Cystoisospora belli, Sarcocystis hominis, and Sarcocystis suihominis are in the family Sarcocystidae, while Cyclospora cayetanensis is in the family Eimeriidae. Many people are familiar with the old name for Cystoisospora belli, Isospora belli, but life cycle (1) and molecular studies indicate that the parasite is a member of the genus Cystoisospora (2), not Isospora.

DESCRIPTION OF THE AGENTS Life Cycles Cystoisospora belli The life cycle is direct (monoxenous), but evidence exists that it can be facultatively heteroxenous (use two hosts). C. belli oocysts are passed in the feces unsporulated or partially sporulated (Fig. 1C and D and 2A and B). Oocysts generally complete sporulation within 72 h, although sporulation time varies between 24 h and >5 days, depending on temperature and humidity. Sporulated oocysts contain two sporocysts, each with four sporozoites, although Caryospora-like oocysts of C. belli (containing one sporocyst with eight sporozoites) have been reported and can comprise up to 5% of the sporulated oocysts in a sample (3). The prepatent period, the time it takes for unsporulated oocysts to appear in the feces after sporulated oocysts are ingested, is 9 to 17 days (4). The patent period, the time from when oocysts are first excreted in the feces until they can no longer be observed in the feces, is quite variable and depends on the immune status of the infected individual. Oocysts can usually be found for 30 to 50 days in immunocompetent patients, while immunosuppressed patients may continue to shed oocysts

*This chapter contains information presented by David S. Lindsay, Steve J. Upton, and Louis M. Weiss in chapter 138 of the 10th edition of this Manual.

for 6 months or more (5). Recurrence of oocyst shedding is common. This prolonged oocyst shedding in immunosuppressed patients is presumably due to recycling of one or more schizogenous stages or activation of dormant extraintestinal monozoic tissue cysts (Fig. 4). Developmental stages of C. belli have been reported for intestinal biopsy specimens of the duodenum, jejunum, and occasionally ileum, and oocysts can be aspirated directly from the duodenal contents. Intestinal development occurs predominantly in epithelial cells, although schizonts (meronts) are occasionally reported from the lamina propria or submucosa (6). At least two generations of schizonts, as well as macrogametocytes (female sexual stage), microgametocytes (male sexual stage), and unsporulated oocysts, have been observed. C. belli sporozoites/merozoites are capable of traveling extraintestinally and becoming dormant as single-organismcontaining tissue cysts (Fig. 4) in a variety of tissues, including lamina propria, mesenteric lymph nodes, liver, and spleen (7–9). These cysts are commonly termed monozoic tissue cysts. Monozoic tissue cysts in histological sections are thick walled and measure 12 to 22 by 8 to 10 μm, and each contains a single dormant sporozoite/merozoite of about 8 to 10 by 5 μm (7, 8). Presumably, the monozoic tissue cysts are capable of reactivating patent infections once immunity wanes. Monozoic tissue cysts can be present in the lamina propria in the absence of oocysts in stool samples (9). The existence of these monozoic tissue cysts has led to speculation that a paratenic (transport) host may be involved in the life cycle of C. belli (7). Paratenic hosts are known to occur in the Cystoisospora species that infect cats and dogs, and it is probable that they occur in the life cycle of C. belli.

Cystoisospora natalensis The life cycle is unknown but presumably monoxenous (direct). The oocysts are smaller and more spherical than those of C. belli (Table 1). Oocysts are passed in the stool unsporulated. At ambient temperature, oocysts can complete sporulation within 24 h (10). The prepatent and patent periods of C. natalensis are unknown. One individual passed unsporulated oocysts for at least 4 days (11). The validity of this species is questionable because oocysts of C. natalensis have been reported only for patients from South Africa and no reports of C. natalensis appear in the literature after 1955 (10, 11).

doi:10.1128/9781555817381.ch141

2425

2426 n

PARASITOLOGY

FIGURE 3 Sporocyst of a Sarcocystis species in a stool sample viewed using differential interference contrast microscopy (A) and autofluorescence using UV light (B). Bar, 10 μm. Courtesy of Alice E. Houk, Department of Biomedical Sciences and Pathobiology, Virginia Tech, Blacksburg, VA. doi:10.1128/9781555817381.ch141.f3

FIGURE 1 Line drawings of unsporulated and sporulated oocysts of Cyclospora cayetanensis (A and B) and Cystoisospora belli (C and D) and a sporulated oocyst (E) and sporocyst (F) of a Sarcocystis species from humans. Bar, 10 μm. doi:10.1128/9781555817381.ch141.f1

Cyclospora cayetanensis The life cycle is monoxenous and involves only humans as hosts. Oocysts are passed in the stool unsporulated (Fig. 1A and 2C). At room temperature (23 to 25°C), small numbers of oocysts may sporulate within 10 to 12 days (3, 12) (Fig. 1B). However, many oocysts require 3 to 4 weeks for

sporozoites to fully develop. Sporulated C. cayetanensis oocysts contain two sporocysts, each with two sporozoites. A structure termed a Stieda body is present in the end of each sporocyst. There are no Stieda bodies in the oocysts of C. belli or Sarcocystis species (Fig. 1). The precise prepatent period is not yet known. However, the onset of clinical signs following infection generally averages 7 to 8 days postinfection and lasts 2 to 3 weeks, but this may range from 1 to >100 days. The length of time that oocysts are shed in the feces is highly variable. Oocysts may be shed in the feces for anywhere from 7 days to several months. Relapse of diarrhea can occur in up to 25% of infected individuals (13). Indigenous infections are confined primarily to tropical, subtropical, or warm temperate regions of the world. Outbreaks occur in other areas of the world due to contaminated foodstuffs obtained from regions of endemicity (85). Developmental stages of C. cayetanensis generally occur within epithelial cells of the lower duodenum and jejunum (14–16). There are two asexual generations followed by sexual stages and oocysts. Stages develop in a supranuclear location within enterocytes (16). An experimental attempt to infect seven healthy human volunteers by oral adminis-

FIGURE 2 Modified Kinyoun’s acid-fast-stained smears demonstrating a Cystoisospora belli oocyst with a sporont (A), a C. belli oocyst with two sporoblasts (B), and a Cyclospora cayetanensis oocyst with a sporont (C). Bar, 1µm. doi:10.1128/9781555817381.ch142.f2

141. Cystoisospora, Cyclospora, and Sarcocystis

n

2427

pria. The oocysts are Cystoisospora-like and contain two sporocysts, each with four sporozoites. The oocyst wall often ruptures as the oocyst makes its way to the intestinal lumen. This results in the shedding of individual sporocysts in the feces (Fig. 3). Individual sporocysts contain four sporozoites. Both oocysts with two sporocysts and individual sporocysts can be seen in the feces of humans with intestinal Sarcocystis infection (Fig. 1E and F). Oocysts and sporocysts are fully sporulated when passed in the feces. For human volunteers, the prepatent period has been reported to be 8 to 39 days, and patent infections can last as long as 18 months. S. hominis occurs on all continents, anywhere cattle or buffaloes have access to human feces and humans ingest raw or undercooked beef. FIGURE 4 Hematoxylin and eosin-stained tissue section demonstrating several monozoic tissue cysts (arrows) of Cystoisospora belli. Note the thick wall that surrounds each single zoite. Bar, 10 μm. doi:10.1128/9781555817381.ch141.f4

tration of microscopically confirmed sporulated oocysts of C. cayetanensis was not successful, as none of the patients developed clinical signs or shed oocysts in their stools (17).

Sarcocystis hominis The life cycle is heteroxenous. Humans are definitive hosts for S. hominis, and bovids are the intermediate hosts. Infection occurs when raw or undercooked meat containing sarcocysts is ingested. Known intermediate hosts include cattle (Bos taurus), American bison (Bison bison), water buffaloes (Bubalus bubalis), and wisents (European bison; Bison bonasus). These intermediate hosts harbor sarcocysts (muscle cysts) that are infective when ingested by humans. Infective bradyzoites (dormant merozoite-like stages) are present in the sarcocysts. The bradyzoites penetrate the human intestinal epithelium and develop as sexual stages (macrogametocytes and microgametocytes) in cells in the lamina propria of the intestine. Fertilization occurs, and the oocysts sporulate in the lamina pro-

Sarcocystis suihominis The life cycle is similar to that described above for S. hominis, except that pigs are the intermediate hosts. The prepatent period is 9 to 10 days, and patency is in excess of 36 days. S. suihominis presumably occurs on all continents, anywhere swine have access to human feces and humans ingest raw or undercooked pork.

Human Muscular Sarcocystis Infection Sarcocysts have been reported as incidental findings from tissue sections of both skeletal and cardiac muscle of nearly 100 humans worldwide. Humans become infected after ingesting Sarcocystis species sporocysts in contaminated food or water. One to two generations of precystic schizogony presumably occur in endothelial cells in capillaries throughout the body. The final generation of merozoites penetrates striated muscle cells and transforms into metrocytes. Metrocytes divide by endodyogeny to produce bradyzoites within the sarcocyst. Clinical signs probably arise from schizogony occurring in endothelial cells of capillaries and the host reaction to developing sarcocysts in muscles (i.e., eosinophilic myositis). There are at least seven distinct types of sarcocysts (18) present in human muscles. This suggests that as many as seven definitive hosts may be able to produce sporocysts infective for humans. Many of these sarcocysts appear similar to sarcocysts found in nonhuman primates.

EPIDEMIOLOGY, TRANSMISSION, AND PREVENTION Cystoisospora belli

FIGURE 5 Sarcocyst (arrow) of a Sarcocystis species in a skeletal muscle biopsy specimen from a male Dutch patient obtained during an outbreak of muscular sarcocystosis (49) among visitors to Tioman Island off the east coast of Malaysia. A sarcocyst wall (arrowhead) surrounds hundreds of bradyzoites. Note the lack of inflammatory response. The patient’s traveling partner was also confirmed to be positive by muscle biopsy. Bar, 10 μm. Courtesy of Douglas H. Esposito and Clifton Drew, National Center for Emerging and Zoonotic Infectious Diseases, CDC, Atlanta, GA. doi:10.1128/9781555817381.ch141.f5

C. belli is found primarily in tropical, subtropical, and warm temperate regions, but reports of indigenous infections have been published from temperate areas as well. Most cases of infection in temperate areas involve foreign travel or homosexual contact. Transmission is via ingestion of sporulated oocysts and possibly the ingestion of raw or undercooked tissues from unknown paratenic hosts. An outbreak of C. belli infections involving ~90 patients was reported in the city of Antofagasta, Chile, in 1977 (19). It was associated with ingestion of vegetables contaminated with irrigation water from a sewage treatment plant (19). Improving sanitation and water quality in areas of endemicity will decrease transmission of C. belli.

Cyclospora cayetanensis Cyclospora is endemic in Central and South America, the Caribbean, Mexico, Indonesia, Asia, Nepal, Africa, India, southern Europe, and the Middle East. In areas of endemicity, there is an increased risk of Cyclospora infection with contact with soil (20) and water (21). Three outbreaks of

2428 n

PARASITOLOGY TABLE 1 Structural data for Cystoisospora, Cyclospora, and Sarcocystis oocysts and sporocysts found in stool samples from humans Mean size (range) (μm)a Species Oocysts Cystoisospora belli Cystoisospora natalensis Cyclospora cayetanensis Sarcocystis hominis Sarcocystis suihominis

Sporocysts

32 × 14 (20–36 × 10–19) Not given (25–30 × 21–24) 9 × 9 (8–10 × 8–10) 19 × 15 (not given) 19 × 13 (19–20 × 12–15)

14 × 10 (12–17 × 7–11) 17 × 12 (not given) 6 × 4 (6–7 × 3–4) 15 × 9 (13–17 × 8–11) 14 × 11 (12–14 × 10–11)

a

Data have been rounded to the nearest micrometer.

waterborne C. cayetanensis infections were reported in a study of waterborne protozoal disease outbreaks from 2004 and 2010 (22). Infections in most temperate areas are correlated with the consumption of imported, contaminated fruits and vegetables, such as basil, raspberries, lettuce, mesclun, and snow peas. Two large outbreaks of cyclosporiasis occurred during the summer of 2013 (85). One was concentrated in Iowa (153 cases) and Nebraska (86 cases) and linked to a restaurant-associated salad mix that contained iceberg lettuce, romaine lettuce, red cabbage, and carrots (85). The second was associated with fresh cilantro (85) and was associated with a restaurant (22 of 30 patrons) in a large outbreak in Texas (278 cases). Individuals in areas of endemicity should wear gloves when gardening to prevent exposure to oocysts of C. cayetanensis. Better washing of produce may help to remove Cyclospora oocysts, but many fruits are delicate. Most of the produce items implicated as transmitting Cyclospora are consumed raw, which does not lend itself to prevention by thermal means. Nonthermal treatments such as high hydrostatic pressure (23) have been shown to inactivate Toxoplasma gondii oocysts, and these methods may be effective in inactivating Cyclospora on produce.

Sarcocystis Species Human intestinal Sarcocystis species are potentially present in any region in the world where cattle, buffaloes, and swine have access to human feces and the life cycle can be maintained. The cycle has not been detected in the United States. Cultural habits that include ingestion of raw meat or undercooked meat products help to maintain this life cycle in areas where Sarcocystis species are endemic. Cooking meat to an internal temperature of >67°C kills T. gondii tissue cysts in the meat (24), and this temperature should also kill tissue cysts of human-infective Sarcocystis species in meat products. Preventing cattle, buffaloes, and swine from consuming human feces will also break the cycle in areas of endemicity. Most cases of human muscular Sarcocystis infection (Fig. 5) have been reported from the Far East, particularly Malaysia (25–27). One study of 100 consecutive autopsy cases from Malaysia found that 21% of tongue sections were positive for sarcocysts (27). This is probably an underrepresentation of the true prevalence because only a small amount of muscle can be examined histologically. Humans become infected by ingesting sporocysts in contaminated water (25) or food.

CLINICAL SIGNIFICANCE Cystoisospora belli C. belli can cause serious and sometimes fatal disease in immunocompetent individuals. Symptoms of C. belli infec-

tion include diarrhea, steatorrhea, headache, fever, malaise, abdominal pain, vomiting, dehydration, and weight loss (6, 12, 28, 29). Blood is not usually present in the feces. Eosinophilia is observed in many patients (30, 31). The disease is often chronic, with parasites present in the feces or biopsy specimens for several months to years. Recurrences are common and can occur as long as 10 years after successful treatment (32). Disease is more severe in infants and young children. Clinical disease from C. belli infection is usually more severe in immunocompromised patients than in immunocompetent patients. C. belli infection produces diarrhea in AIDS patients that is often very fluid and secretory-like and leads to dehydration requiring hospitalization (30). Fever and weight loss are also common findings. Other opportunistic pathogens are also common copathogens in these patients. C. belli superinfection of the small bowel was seen in a patient who was immunosuppressed with systemic corticosteroids to aid in treatment of eosinophilic gastroenteritis (31). C. belli has been observed in both renal transplant (33) and liver transplant (34) patients. C. belli-induced intestinal lesions and responses to chemotherapy are usually similar to those observed in immunocompetent patients. C. belli has been observed in patients with concurrent Hodgkin’s disease (6), non-Hodgkin’s lymphoproliferative disease (35), human T-cell leukemia virus type 1-associated adult T-cell leukemia (36), and acute lymphoblastic leukemia and human T-cell leukemia virus type 1-associated T-cell lymphoma (37). These patients respond to specific anti-C. belli treatment. Extraintestinal cyst-like stages have been documented for AIDS patients and may play a role in relapse of infection (7). These usually contain a single merozoite-like stage (Fig. 4) and are called monozoic tissue cysts. Many thousands of these stages can be present (7). Infections with C. belli in the gallbladder epithelium (38–40) and endometrial epithelium (41) have been reported, and oocysts have been observed in bile samples (42). Clinical signs in patients with parasites in these locations are not specific for coccidiosis, and parasites are located after tissue biopsy as part of a diagnostic workup. Infection of the biliary tract with C. belli was the first HIV-related opportunistic infection in one patient, and it may have represented an AIDS-defining infection in that case (40). The parasites probably reach these extraintestinal sites as merozoites from the gut or zoites from extraintestinal locations, and the epithelial cells of these tissues are permissive to parasite entrance and multiplication.

Cyclospora cayetanensis Nonbloody watery diarrhea is the main clinical symptom of C. cayetanensis infection. Symptoms of nausea, fatigue, abdominal cramps, and fever were reported in >50% of clinical cases in one foodborne outbreak, with headache

141. Cystoisospora, Cyclospora, and Sarcocystis n 2429

and vomiting occurring in 45 to 30% of these patients (13). Some individuals can be infected and show no clinical signs. In most immunocompetent patients, typical symptoms of cyclosporiasis include cycles of diarrhea with anorexia, malaise, nausea, and cramping and periods of apparent remission. C. cayetanensis infection can be associated with biliary disease in both immunosuppressed patients and immunologically normal patients (43). Developmental stages of C. cayetanensis have been seen in the gallbladder epithelium of AIDS patients with acalculous cholecystitis (44). Oocysts can be observed in the bile of patients with active biliary disease.

Intestinal Sarcocystis Infections Clinical Sarcocystis infections in humans can manifest primarily as intestinal disease if infected meat is ingested or as muscular disease if sporocysts are ingested (45). Intestinal disease occurs soon after consumption of infected meat (3 to 6 h) and is characterized by nausea, abdominal pain, and diarrhea. Intestinal disease can be more severe in individuals who have additional enteropathogens present in the gut. Intestinal Sarcocystis infection combined with invasion by Gram-positive bacteria has been associated with several cases of segmental enterocolitis in Thailand (46). Experimental studies with human volunteers have produced moresevere disease in those who have ingested pork containing S. suihominis than in those who have ingested beef containing S. hominis (45). Some individuals can be infected and show no clinical signs.

Muscular Sarcocystis Infections Muscular Sarcocystis infections (Fig. 5) in humans are usually subclinical or associated with only mild clinical signs and are usually considered incidental findings (17, 47, 48). Clinical case reports are from Southeast Asia. Three outbreaks of acute muscular sarcocystosis have been reported from Malaysia (25, 26, 49). In the first reported outbreak, clinical signs associated with muscular Sarcocystis infection occurred in 7 of 15 members of a U.S. combat unit (25). The signs developed about 3 weeks after the troops returned from the jungle and were fever, myalgias, bronchospasm, fleeting pruritic rashes, transient lymphadenopathy, and subcutaneous nodules. Eosinophilia, elevated erythrocyte sedimentation rate, and elevated levels of muscle creatine kinase were present in these troops (25). The second (26) and third (49) outbreaks occurred in travelers returning from Tioman Island off the east coast of Malaysia. The second outbreak, during the summer of 2011, was in 32 patients, most of whom were from Germany (~50%), other European countries, North America, and Asia. Within days or weeks of returning home, most patients experienced fever and muscle pain (26). All had peripheral eosinophilia, and most had elevated serum creatinine phosphokinase levels (26). The third outbreak was reported in 65 patients returning from Tioman Island (26) and occurred during July and August of 2011 and 2012. The 65 patients originated from Germany (n = 25), France (n = 20), the Netherlands (n = 12), Switzerland (n = 3), Belgium (n = 2), Spain (n = 2), and Singapore (n = 1) (49). Most patients experienced fever and myalgia, while fewer had arthralgia, asthenia, headache, cough, and diarrhea (49). Decreased immunity or cancer may make individuals more susceptible to muscular Sarcocystis infection. Sarcocysts have been reported in the heart of a patient with Hodgkin’s disease (50) and the larynx of a patient with squamous cell carcinoma (51).

COLLECTION, TRANSPORT, AND STORAGE OF SPECIMENS The results obtained in the diagnostic laboratory are only as good as the material presented for testing. Choosing the appropriate sample and sample fixative is extremely important (52). Universal precautions should be followed when fresh stool samples are handled. If samples are to be sent to another laboratory for diagnosis, they should be fixed in an appropriate fixative. A 5% or 10% formalin solution is an appropriate fixative for stools suspected of containing intestinal coccidia. Formalin fixation does not interfere with some of the immunodetection methods currently employed to detect Cryptosporidium and Giardia duodenalis, which is a drawback of polyvinyl alcohol fixative. Oocyst structure lasts for several months when stools are stored at 4°C in formalin fixatives.

DIRECT EXAMINATION Microscopy Oocysts of C. belli and C. cayetanensis and sporocysts of Sarcocystis species are readily identified in fresh unstained wet mounts, based on their characteristic sizes and morphologies (Table 1; Fig. 1). This is especially true if oocysts and sporocysts are present in large numbers. Autofluorescence of oocysts of C. belli and C. cayetanensis and oocysts and sporocysts of Sarcocystis (Fig. 3) is an especially useful tool (53, 54) and has replaced many of the staining techniques previously used for these parasites in laboratories equipped with appropriate fluorescent microscopes. Concentration techniques, such as formalin-ethyl acetate (rarely formalin-ether) sedimentation or sucrose centrifugal flotation, are helpful when few oocysts are present. Sucrose centrifugal flotation has been found to be superior to formalin-ether sedimentation for demonstrating oocysts of C. cayetanensis (55). The same should be true for oocysts of C. belli and sporocysts of Sarcocystis spp. Few laboratories employ sucrose concentration, and fortunately, direct wet smears can be very useful: their utility approaches that of sucrose centrifugal flotation when coupled with autofluorescence examination (55). Staining procedures may adversely affect the autofluorescence of oocysts and sporocysts. Stained fecal smears have been widely used to demonstrate C. cayetanensis oocysts and, to a lesser extent, C. belli oocysts (53, 56). C. belli and C. cayetanensis oocysts stain red with the modified Kinyoun’s acid-fast stain, and this method is widely used (Fig. 2). The main drawbacks are that staining can be variable and some oocysts do not stain (57). Oocysts usually do not stain with trichrome, chromotrope, or Gram-chromotrope stain (57). Some C. cayetanensis oocysts stain light blue with Giemsa stain (57). Variations on the safranin staining technique stain C. cayetanensis oocysts orange or pinkish orange, and heating and other treatments have been used to increase the staining frequency of oocysts. Flow cytometry has been used to detect C. cayetanensis oocysts in human stool samples (58). The results of flow cytometry examination were similar to those of microscopy, and preparation times for the two methods were similar (58). A single negative stool specimen is not conclusive in the examination of stools for coccidial parasites; a total of three or more stool specimens collected on subsequent days need to be examined before coccidial infection can be ruled out. Liquid stool samples can be concentrated by centrifugation, and the pellet may be used for examination by use of wet mounts, concentration techniques, or stained smears.

2430 n

PARASITOLOGY

Large numbers of oocysts may make diagnosis less challenging but do not always translate directly to the severity of clinical signs. Some individuals may excrete oocysts and be asymptomatic. Cases of muscular Sarcocystis infection are diagnosed based on the detection of sarcocysts in muscle samples taken from biopsy specimens or postmortem samples.

Culture In vitro culture of intestinal coccidial parasites is most often used as a tool to study developmental biology or to identify active chemotherapeutic agents. It presently has limited use in diagnosis of active human infection. Only minimal development of C. belli occurs in human (ileocecal adenocarcinoma [HCT-8 cells] or epithelial carcinoma of the lung [A549]) or mammalian (African green monkey kidney [Vero] or Madin-Darby bovine kidney) cell cultures (59). No reports on development of C. cayetanensis in cell culture have been published. Bradyzoites of S. suihominis undergo sexual development and produce oocysts in cell culture (60). Vascular schizont stages of human Sarcocystis species have not been reported for in vitro systems, but continuous cultures of several mammalian and avian species have been reported.

Antigen Detection The inability to produce stages in cell cultures and provide a source of diagnostic antigens has limited the usefulness of antigen detection for these coccidial parasites of humans.

Nucleic Acid Detection There are no U.S. Food and Drug Administration-approved nucleic acid tests for the detection of infections with C. belli, C. cayetanensis, or Sarcocystis species. Several research laboratories have developed nucleic acid-based detection tests to demonstrate infection with these parasites in stool and tissue samples. Detection of C. belli by PCR with primers based on internal transcribed spacer (ITS) region and small-subunit RNA sequences has been reported (2, 61, 62). Three different genotypes of C. belli can be identified with PCR and restriction fragment length polymorphism using MboII digestion (62) Coinfection of a single patient with two different genotypes has been observed (62). Molecular diagnosis of cystoisosporiasis employing extended-range PCR screening has been proven to detect C. belli in biopsy material from a patient who was fecal examination and histological examination negative on initial testing (63). Developmental stages were eventually observed in biopsy material and confirmed the findings (63). A real-time PCR using the internal transcribed spacer region 2 small-subunit RNA sequences has been developed to detect C. belli in stool samples (61). Much attention has been placed on molecular methods to detect C. cayetanensis oocysts in stools, in water samples, and on produce because of the numerous outbreaks of C. cayetanensis infections (64). The 18S rRNA gene is presently the most frequently used target. Because C. cayetanensis is closely related to Eimeria species (65) from vertebrates, it is important that tests designed to detect C. cayetanensis in the water or on produce be examined for cross-reactivity to Eimeria spp. (66). Cyclospora species infecting mammals other than humans may also be present in water samples or on produce, and proofs of specificity are needed for these tests designed to look at environmental sources of C. cayetanensis and to detect C. cayetanensis oocysts on produce. Quantitative PCR assays have been developed for C. cayetanensis oocysts in stool samples (67). This method de-

tected DNA of the 18S ribosomal gene sequence from as little as 1 oocyst of C. cayetanensis per 5 ml of reaction volume. A multiplex PCR to detect Cyclospora, Cystoisospora, and microsporidia in stool samples has been developed (68). S. hominis sporocysts in human stool can be detected using PCR-restriction fragment length polymorphism and sequencing of a partial 18S rRNA gene product. Several PCR methods have been developed to detect sarcocysts in the muscles of animals, and the U.S. Centers for Disease Control and Prevention (CDC) is presently attempting to develop a Sarcocystis-specific PCR to detect muscular Sarcocystis infection (26).

SEROLOGIC TESTS The inability to obtain usable quantities of antigens from C. belli, C. cayetanensis, and Sarcocystis spp. has greatly limited the use of serologic diagnostic tests for these parasites in human stool samples. It is very difficult to obtain enough oocysts or sporocysts from feces to conduct serologic tests. The development of an enzyme-linked immunosorbent assay that measures IgG and IgM antibodies, using C. cayetanensis oocysts as antigen, has been reported (69). This study is questionable because the authors stated that they obtained their oocysts from experimentally infected guinea pigs, yet well-controlled studies indicate that humans are the only suitable host for C. cayetanensis (70). Attempts were made to develop an indirect fluorescent-antibody assay to detect IgG, IgA, and IgM specific for C. cayetanensis, using sectioned oocysts (71). None of the sera from four patients with positive stools reacted in the indirect fluorescent-antibody assay. The CDC is presently attempting to develop a Sarcocystis-specific Western blot test to detect muscular Sarcocystis infection (49; D. H. Esposito and S. Handali, personal communication, April 2013). As of this writing, no test is available.

TREATMENT Cystoisospora belli The drug of choice for the treatment of C. belli is trimethoprim-sulfamethoxazole. A dose of trimethoprim (160 mg)sulfamethoxazole (800 mg) two to four times a day for 10 to 14 days results in clearance of parasites, a decrease in diarrhea, and a decrease in abdominal pain within a mean of 2.5 days after treatment (72). Before the advent of combination antiretroviral therapy (cART), it was recommended that patients with HIV-1 infection and CD4+ cell counts of 20 established Cryptosporidium species, including C. hominis in humans; C. parvum in humans and preweaned ruminants; C. viatorum in humans; C. andersoni, C. bovis, and C. ryanae in weaned calves and adult cattle; C. xiaoi and C. ubiquitum in sheep and goats; C. suis and C. scrofarum in pigs; C. canis in dogs; C. felis in cats; C. cuniculus in rabbits; C. wrairi in guinea pigs; C. muris and C. tyzzeri in rodents; C. fayeri and C. macropodum in marsupials; C. meleagridis, C. baileyi, and C. galli in birds; C. varanii and C. serpentis in reptiles; C. fragile in amphibians; and C. molnari and C. scophthalmi in fish. There are also >40 host-adapted Cryptosporidium genotypes that do not yet have species names, such as Cryptosporidium horse, skunk, and hedgehog genotypes (8–12). These species and genotypes biologically, morphologically, and phylogenetically belong to three groups: intestinal, gastric, and piscine species and genotypes (13). Most of these Cryptosporidium species and genotypes have not been found in humans.

DESCRIPTION OF THE AGENT Cryptosporidium spp. are intracellular parasites that primarily infect epithelial cells of the intestine and biliary ducts. In severely immunosuppressed persons, the respiratory tract is sometimes involved. The involvement of the respiratory system in immunocompetent persons may be more common than previously believed, although its role in cryptosporidiosis transmission is not yet clear (14). The infection site varies according to species, but almost the entire development of Cryptosporidium spp. occurs between the two lipoprotein layers of the membrane of the epithelial cells. Cryptosporidium infections in humans or other susceptible hosts start with the ingestion of viable oocysts (Fig. 1). Upon contact with gastric and duodenal fluid, four sporozoites are liberated from each excysted oocyst; invade the epithelial cells; develop to trophozoites surrounded by a parasitophorous vacuole; and undergo two or three generations of asexual multiplication and one generation of sexual reproduction, leading to the formation of new oocysts. The latter are sporulated in situ, excreted into the environment with feces, and can initiate infection in a new host upon ingestion without further development (Fig. 1). The time from ingestion of infective oocysts to the completion of endogenous development and excretion of new oocysts varies with species, hosts, and infection doses; it is usually between 4 and 10 days (15, 16). In addition to the classic coccidian developmental stages, a gregarine-like extracellular stage was described in C. parvum, which supposedly can go through multiplication via syzygy, a sexual reproduction process involving the end-toend fusion of two or more parasites (7). Currently, nearly 20 Cryptosporidium species and genotypes shave been reported in humans, including C. hominis, C. parvum, C. meleagridis, C. felis, C. canis, C. cuniculus, C. ubiquitum, C. viatorum, C. muris, C. suis, C. fayeri, C. andersoni, C. bovis, C. scrofarum, C. tyzzeri, and Cryptosporidium horse, skunk, hedgehog, and chipmunk I genotypes. Humans are most frequently infected with C. hominis and C. parvum. The former almost exclusively infects humans and nonhuman primates and thus is considered an anthroponotic parasite, whereas the latter mostly infects humans and ruminants and thus is considered a zoonotic pathogen. Other species, such as C. meleagridis, C. felis, C. canis, C. cuniculus, C. ubiquitum, and C. viatorum, are less common. The remaining Cryptosporidium species and genotypes have been found only in a few human cases (17, 18). These Cryptosporidium spp. infect both immunocompetent and immunocom-

doi:10.1128/9781555817381.ch142

2435

2436 n

PARASITOLOGY

FIGURE 1 Life cycle of Cryptosporidium spp. Sporulated oocysts, containing four sporozoites, are excreted by the infected host through feces and possibly other routes such as respiratory secretions (1). Transmission of Cryptosporidium spp. in humans occurs mainly through contact with infected persons (for C. hominis and C. parvum) or animals (for C. parvum mostly) and consumption of contaminated water and food (2). Following ingestion (and possibly inhalation) by a suitable host (3), excystation (a) occurs. The sporozoites are released and parasitize epithelial cells (b, c) of the gastrointestinal tract or other tissues such as the respiratory tract. In these cells, the parasites undergo asexual multiplication (schizogony or merogony) (d to f) and then sexual multiplication (gametogony), producing microgamonts (male) (g) and macrogamonts (female) (h). Upon fertilization of the macrogamonts by the microgametes (i), oocysts (j, k) develop that sporulate in the infected host. Two different types of oocysts are produced, the thick-walled oocyst (j), which is commonly excreted from the host; and the thin-walled oocyst (k), which is primarily involved in autoinfection. Oocysts are infective upon excretion, thus permitting direct and immediate fecal-oral transmission. Courtesy of DPDx (http://www.cdc.gov/dpdx/). doi:10.1128/9781555817381.ch142.f1

promised persons. The distribution of these species in humans varies among different geographic areas and socioeconomic conditions, with C. canis and C. felis mostly seen in humans in developing countries, C. ubiquitum mostly in industrialized nations, and C. cuniculus mostly in the United Kingdom. This is probably the result of differences in infection sources and transmission routes (17).

EPIDEMIOLOGY, TRANSMISSION, AND PREVENTION Cryptosporidium spp. have a worldwide distribution, and their oocysts are ubiquitous in the environment. In the United States, the number of annual reported cases of cryptosporidiosis has increased more than 2-fold since 2004; there were 2,769 to 3,787 annual reported cases between

142. Cryptosporidium

1999 and 2002, 3,505 to 8,269 between 2003 and 2005, 6,479 to 11,657 between 2006 and 2008, and 7,656 to 8,951 between 2009 and 2010 (19–22). Currently, it is estimated that there are ~750,000 cases of cryptosporidiosis in the United States each year, as 98.6% of cases remain undiagnosed or unreported (5). The estimated annual cost of hospitalization alone due to cryptosporidiosis in the United States is $45 million, with a per-case cost of $16,797 (23). Humans can acquire cryptosporidiosis through several transmission routes, such as direct contact with infected persons or animals and consumption of contaminated water (drinking or recreational) or food (1, 17). However, the relative role of each in the transmission of Cryptosporidium infection in humans is unclear.

Susceptible Populations In developing countries, human Cryptosporidium infection occurs mostly in children younger than 2 years and is one of the top five causes of diarrhea in this age group (2, 3, 16). In industrialized nations, pediatric cryptosporidiosis occurs in children later than in developing countries, probably due to delayed exposures to contaminated environments as a result of better hygiene (19, 21, 22, 24, 25). Cryptosporidiosis is also common in elderly people in nursing homes, where person-to-person transmission probably plays a major role in the spread of Cryptosporidium infections (26). In the general population, a substantial number of adults are susceptible to Cryptosporidium infection, as sporadic infections occur in all age groups in the United States and the United Kingdom, and travel to developing countries and consumption of contaminated food or water can frequently lead to infection (27–30). Cryptosporidiosis is common in immunocompromised persons, including AIDS patients, persons with primary immunodeficiency, and cancer and transplant patients undergoing immunosuppressive therapy (1, 31–37). Hemodialysis patients with chronic renal failure and renal transplant patients commonly develop cryptosporidiosis (34, 38–41). In HIV-positive persons, the occurrence of cryptosporidiosis increases as the CD4+ lymphocyte cell counts fall, especially below 200 cells/μl (32).

Anthroponotic versus Zoonotic Transmission Studies in the United States and Europe have shown that cryptosporidiosis is more common among homosexual men than among persons in other HIV transmission categories (42), indicating that direct person-to-person or anthroponotic transmission of cryptosporidiosis is common. Contact with persons with diarrhea has been identified as a major risk factor for sporadic cryptosporidiosis in industrialized countries (28, 43–46). This is exemplified by the high prevalence of cryptosporidiosis in day care facilities and nursing homes and among mothers with young children in these countries. Only a few case-control studies have assessed the role of zoonotic transmission in the acquisition of cryptosporidiosis in humans. In industrialized countries, contact with farm animals (especially cattle) is a major risk factor in sporadic cases of human cryptosporidiosis (28, 30, 43, 44, 47, 48). Contact with pigs, dogs, or cats was a risk factor for cryptosporidiosis in children in Guinea-Bissau and Indonesia in one study (49, 50). A weak association was observed between the occurrence of cryptosporidiosis in HIV-positive persons and contact with dogs in another study (51). In other studies, no increased risk in the acquisition of cryptosporidiosis was associated with contact with companion animals (52). The distribution of C. parvum and C. hominis in humans is a reflection of the role of different transmission routes.

n

2437

Thus far, studies conducted in developing countries have shown a predominance of C. hominis in children or HIVpositive adults. This is also true for most areas in the United States, Canada, Australia, and Japan. In Europe and New Zealand, however, several studies have shown almost equal prevalence of C. parvum and C. hominis in both immunocompetent and immunocompromised persons (17). In contrast, children in the Middle East are mostly infected with C. parvum (53). The differences in the distribution of Cryptosporidium genotypes in humans are considered to be an indication of differences in infection sources (17, 48, 54, 55); the occurrence of C. hominis in humans is most likely due to anthroponotic transmission, whereas C. parvum in a population can be the result of both anthroponotic and zoonotic transmission. Thus, in most developing countries, it is possible that anthroponotic transmission of Cryptosporidium plays a major role in human cryptosporidiosis, whereas in Europe, New Zealand, and rural areas of the United States, both anthroponotic and zoonotic transmissions are important. Recent subtyping studies based on sequence analyses of the 60-kDa glycoprotein (gp60) gene have shown that many C. parvum infections in humans are not results of zoonotic transmission (17). Among several C. parvum subtype families identified, IIa and IIc are the two most common families. The former has been identified in both humans and ruminants and thus can be a zoonotic pathogen, whereas the latter has been seen only in humans (13, 17, 56) and thus is an anthroponotic pathogen. In developing countries, most C. parvum infections in children and HIV-positive persons are caused by the subtype family IIc, with IIa largely absent, indicating that anthroponotic transmission of C. parvum is common in these areas (13, 17). In contrast, both IIa and IIc subtype families are seen in humans in developed countries. Even in the United Kingdom, where zoonotic transmission is known to play a significant role in the transmission of human cryptosporidiosis, anthroponotic transmission of C. parvum also occurs (57). Another C. parvum subtype family commonly found in sheep and goats, IId, is the dominant C. parvum subtype family in humans in Middle Eastern countries (53). Results of multilocus subtyping support the conclusions of gp60 subtyping studies (58–61).

Waterborne Transmission Epidemiologic studies have frequently identified water as a major route of Cryptosporidium transmission in areas where the disease is endemic (62). In most tropical countries, Cryptosporidium infections in children usually peak during the rainy season; thus, waterborne transmission probably plays a role in the transmission of cryptosporidiosis in these areas (63–65). Seasonal variations in the incidence of human Cryptosporidium infection in industrialized nations have also been partially attributed to waterborne transmission (27, 28, 30, 66). In the United States, there is a late summer peak in sporadic cases of cryptosporidiosis (22, 28, 30), which is largely due to increased participation in recreational activities such as swimming and water sports (67). In a Canadian study, swimming in a lake or river was identified as a risk factor (45). The role of drinking water in sporadic Cryptosporidium infection is not clear. In Mexican children living near the U.S. border, cryptosporidiosis is associated with consumption of municipal water instead of bottled water (68). In England, there is an association between the number of glasses of tap water drunk at home each day and the occurrence of sporadic cryptosporidiosis (43). In the United States, drinking untreated surface water was identified as a

2438 n

PARASITOLOGY

risk factor for the acquisition of Cryptosporidium in casecontrol studies (46, 69), and residents living in cities with surface-derived drinking water generally have higher antibody levels against Cryptosporidium in their blood than those living in cities with groundwater as drinking water, indicating that drinking water plays a role in the transmission of human cryptosporidiosis (70). Nevertheless, case-control studies conducted with both immunocompetent persons and AIDS patients in the United States and Canada have failed to show a direct linkage of Cryptosporidium infection to drinking water (71–73). Numerous waterborne outbreaks of cryptosporidiosis have occurred in the United States, Canada, the United Kingdom, France, Australia, Japan, and other industrialized nations (4, 74). These include outbreaks associated with both drinking water and recreational water (swimming pools and water parks). After the massive cryptosporidiosis outbreak in Milwaukee, WI, in 1993, the water industry has adopted more-stringent treatments of source water. Currently, the number of drinking-water-associated outbreaks is in decline in the United States and the United Kingdom, and most outbreaks in the United States are associated with recreational water (30, 67). Even though five Cryptosporidium species are commonly found in humans, C. parvum and C. hominis are responsible for most cryptosporidiosis outbreaks, with C. hominis responsible for more outbreaks than C. parvum (17). This is even the case for the United Kingdom, where C. parvum and C. hominis are both common in the general population. Recently, there was one drinkingwater-associated cryptosporidiosis outbreak caused by C. cuniculus (75). An outbreak of C. meleagridis also occurred in a high school dormitory in Japan, although the role of waterborne transmission in the occurrence of the outbreak was not clear (76).

Foodborne Transmission Foodborne transmission is also important in cryptosporidiosis epidemiology. Cryptosporidium oocysts have been isolated often from fruits, vegetables, and shellfish (77–80). Direct contamination of food by fecal materials from animals or food handlers has been implicated in several foodborne outbreaks of cryptosporidiosis in industrialized nations. In most instances, human infections were usually due to consumption of contaminated fresh produce and unpasteurized apple cider or milk (81–87). Very few case-control studies have examined the role of contaminated food as a risk factor in the acquisition of Cryptosporidium infection in areas where the disease is endemic. A pediatric study in Brazil failed to show any association between Cryptosporidium infection and diet or type of food hygiene (52). Case-control studies conducted in the United States, the United Kingdom, and Australia have actually shown a lower prevalence of Cryptosporidium infection in immunocompetent persons with frequent consumption of raw vegetables (28, 43, 46, 88), probably because of continuous exposure to Cryptosporidium-contaminated fresh produce. It is estimated that ~8% of Cryptosporidium infections in the United States are foodborne (5).

Prevention As for any pathogens that are transmitted by the fecal-oral route, good hygiene is the key in preventing the acquisition of Cryptosporidium infection (89). Immunosuppressed persons especially should take necessary precautions in preventing the occurrence of cryptosporidiosis (90). This includes washing hands before preparing food and after using the bathroom, changing diapers, and contacting pets or soil

(including gardening); avoiding drinking water from lakes and rivers, swallowing water in recreational activities, and drinking unpasteurized milk, milk products, and juices; and following safe-sex practices (avoiding oral-anal contact). During cryptosporidiosis outbreaks or when a community advisory to boil water is issued, individuals should boil water for 1 min to kill the parasite or use a tap water filter capable of removing particles 90%) and sensitivities (98 to 100%) (105, 122–127). However, sensitivities of 68 to 75% were shown in some studies for some assays (105, 128–130). High false-positive rates (positive predictive value, 56%) for several rapid assays have been reported recently in clinical diagnosis of cryptosporidiosis in the United States (131). This has prompted the Council of State and Territorial Epidemiologists to change the case definition of rapid assay-positive cases from confirmed cases to probable cases. It has also been shown recently that some rapid assay kits have low sensitivity (90% sensitivity and specificity (43, 45).

Treatment The major goals of therapy are to prevent irreversible lesions and to alleviate symptoms. Surgical excision of nodules is recommended when the nodules are located on the head because of the proximity of the microfilaria-producing adult worms to the eye, but chemotherapy is the mainstay of treatment. Ivermectin, a semisynthetic macrocyclic lactone, is now considered the first-line therapy for onchocerciasis. It is characteristically given yearly or semiannually. Most patients have limited or no reaction to treatment. Pruritus,

144. Filarial Nematodes n 2467

cutaneous edema, and/or a maculopapular rash occur in approximately 1 to 10% of treated individuals. Significant ocular complications are extremely rare, as is hypotension (1 in 10,000). In areas of Africa where O. volvulus and L. loa are coendemic, however, ivermectin is contraindicated because of severe posttreatment encephalopathy seen in patients who show heavy L. loa microfilaremia (46). There has been some recent evidence that doxycycline-based therapy targeting the Wolbachia endosymbiont, which is present in Onchocerca but not L. loa, might have improved efficacy in areas of coinfection with no increase in adverse events (47). Ivermectin is also contraindicated for use in pregnant or breastfeeding women, based on toxicity and teratogenicity data from animal studies. Although ivermectin treatment results in a marked drop in microfilarial density, its effect can be short-lived (15 main lateral uterine hooks branches Squared, four suckers, 95% for Taenia, Diphyllobothrium, and Hymenolepis species infections

a

Adapted from reference 1.

Praziquantel levels in serum are therefore lowered when any of these drugs are coadministered. Cimetidine, which inhibits P450-mediated metabolism, can be given concurrently to increase levels of praziquantel in plasma. Levels in plasma peak after 1.5 to 2 h, and after a single 40-mg/ kg dose, peak levels have been reported as 1.007 to 1.625 mg/liter (33). Praziquantel does not cross the blood-brain barrier well, so levels in CSF are only approximately 20 to 25% of those in plasma (14, 32). It is 80% protein bound, and its half-life in serum is 1 to 3 h. It is not dialyzable, and no adjustment in dose is recommended in either renal or hepatic insufficiency. Spectrum of activity. Praziquantel is active against the larval and adult stages of many trematodes. It is the drug of choice for schistosomiasis and is effective for all Schistosoma species that infect humans. It is used both for treatment of individuals and in mass community chemotherapy programs and leads to decreased transmission and prevalence of infection. Praziquantel is also used for treatment of opisthorchiasis, clonorchiasis, paragonimiasis, and intestinal fluke infections, including fasciolopsiasis, heterophyiasis, and metagonimiasis. In contrast to other human trematode infections, praziquantel has not proven to be effective in the treatment of Fasciola hepatica infection. Many cestode infections can also be treated with praziquantel. Most tapeworm infections respond, including those caused by Taenia, Diphyllobothrium, and Hymenolepis species. Because praziquantel does not kill eggs, precautions should be taken to prevent autoinfection and laboratory-acquired infection, particularly for Taenia solium. Praziquantel is also used in the treatment of neurocysticercosis as an alternative or adjunct to albendazole, although its overall benefit remains unclear. Praziquantel has also been used in combination with albendazole for treatment of echinococcal infections. Praziquantel has high protoscolicidal activity in vitro, and some reports have suggested superior efficacy of the combination to that of either drug alone (34–36). Adverse effects. Adverse effects of praziquantel are generally mild, but many studies report that some side effects occur in >30% of patients. Common reactions include dizziness, lethargy, headache, nausea, and abdominal pain. Hypersensitivity reactions occur rarely (37, 38). Severe adverse reactions are uncommon, although administration to individuals with neurocysticercosis can result in seizures and neurological sequelae related to precipitation of an inflammatory response (39). Animal studies do not suggest a teratogenic effect of praziquantel (pregnancy category B), but an increased abor-

tion rate has been seen in rats. There are minimal data on its safety in humans, but there is a very low potential for adverse effects on either the mother or her unborn child (40), and when praziquantel has been used during pregnancy, no increase in abortion rates, preterm deliveries, or congenital abnormalities has been noted (41). Consequently, praziquantel can be given after the first trimester. No adverse effects of praziquantel administration during lactation have been reported, but it is excreted in human breast milk and discontinuation of breast feeding on the day of therapy and for the following 72 h is sometimes suggested. However, owing to available data regarding its safety profile, in 2002 the World Health Organization recommended that it can be considered for use in pregnant and lactating women (42). It is FDA approved for children aged 4 years and over.

Ivermectin Ivermectin is a semisynthetic macrocyclic lactone derivative of avermectins, which are natural substances derived from the actinomycete Streptomyces avermitilis. Major indications for its use are shown in Table 6. It was initially developed as an agent for veterinary use but is now used widely in humans. Ivermectin is a potent oral agent with relatively broad-spectrum anthelmintic activity. It has been approved by the U.S. FDA as an oral therapy for onchocerciasis and uncomplicated strongyloidiasis (1) and as a topical treatment for head lice. Mechanism of action. Ivermectin causes an influx of chloride ions across glutamate-gated chloride channels in nerve and muscle cell membranes, resulting in hyperpolarization of the affected cells and consequent paralysis and death of parasites (4, 43–46). It has also been postulated that ivermectin may act as an antagonist of the neurotransmitter γ-aminobutyric acid (47). Although specific ivermectin binding sites have been identified in mammalian brain tissue, the affinity of ivermectin for sites within parasites is ∼100 times greater than that for mammalian tissue. Pharmacokinetics. Ivermectin is available as an oral preparation. There is no parenteral preparation of ivermectin approved by the FDA, but it has been given via the rectal and subcutaneous routes to critically ill patients with disseminated strongyloidiasis who are unable to tolerate oral therapy (48, 49). It is rapidly absorbed following oral administration. Bioavailability is thought to increase ∼2.5fold if it is taken with a high-fat meal; recommendations are to administer the drug on an empty stomach with water. Ivermectin is metabolized in the liver and excreted almost

2534 n ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS TABLE 6 Major indications for ivermectina Indication Ascaris lumbricoides Cutaneous larva migrans Gnathostoma spinigerum Onchocerca volvulus Strongyloides stercoralis

Trichuris trichiura Ectoparasites: scabies and lice

Usual dose

Reported efficacy

Single dose of 150–200 μg/kg 200 μg/kg daily (usually 12 mg) for 1–2 days 200 μg/kg daily for 2 days Single dose of 150 μg/kg, repeat every 6–12 mo until asymptomatic 200 μg/kg daily for 2 doses (doses given either on consecutive days or 2 weeks apart) 200 μg/kg daily for 3 days 200-μg/kg dose, repeat after 2 wk for scabies

Cure rate of 78–99% Cure rate of 77–100% Cure rate of 76–95% Skin microfilarial counts reduced by 85–95%, and levels remain suppressed by >90% at 1 yr Cure rate of 85–97% in uncomplicated infection (normal or immunocompromised hosts) Cure rate of 35–84% Almost 100% efficacy

a

Adapted from reference 1.

entirely in the feces. Peak levels in serum occur at 4 to 5 h, and levels of ∼46 μg/liter have been reported after a single 12-mg dose (15). It is highly protein bound in plasma, and it has a half-life of 10 to 18 h. The drug accumulates in fat tissue and does not readily cross the blood-brain barrier (50). Spectrum of activity. Ivermectin is the drug of choice for onchocerciasis and strongyloidiasis. In onchocercal infections, ivermectin does not significantly affect the viability of adult worms, but it impairs release of microfilariae, is a potent microfilaricide, and leads to a sustained reduction in microfilaremia for many months (44). It can be used both for the treatment of individual patients and in mass chemotherapy programs in areas where onchocerciasis is endemic. The role of combination therapy with ivermectin is being explored. In uncomplicated strongyloidiasis, ivermectin has excellent efficacy in immunocompetent patients (47). In disseminated strongyloidiasis, ivermectin is administered daily until stool and sputum exams are negative for larvae. Ivermectin also has activity against microfilariae of Wuchereria bancrofti, Brugia malayi, and Loa loa. It does not have a significant effect on adult worm viability in these infections, so reduced microfilaremia is sustained only with repeated doses, and it has not replaced DEC as first-line therapy for these infections. Ivermectin also has activity against Mansonella ozzardi and Mansonella streptocerca microfilariae. Ivermectin also has efficacy against many intestinal helminths, including Ascaris lumbricoides, T. trichiura, Enterobius vermicularis, and cutaneous larva migrans (45, 51–53). However, it is not generally used as first-line treatment for these indications due to the widespread availability and excellent efficacy of albendazole. It also can be used for treatment of gnathostomiasis (54, 55). It is not active against trematodes or cestodes. In addition to its anthelmintic activity, ivermectin is the drug of choice for ectoparasitic infestations, including those of scabies and lice. Adverse effects. Ivermectin is generally well tolerated. Most of the adverse effects that occur following its administration are a result of the host’s immune response to destruction of parasites rather than to toxic effects of the drug per se. Adverse effects include fever, rash, dizziness, pruritis, myalgia, arthralgia, and tender lymphadenopathy; the severity of these symptoms relates to the pretreatment intensity of infection rather than to ivermectin concentrations in serum (56). Transaminitis is occasionally reported. Severe

reactions occasionally occur, including a hypersensitivity response to dying microfilarial parasites known as the Mazzotti reaction. This anaphylactoid response is characterized by allergic manifestations including pruritis, edema, fever, and systemic hypotension. However, these reactions are primarily restricted to individuals with high parasite loads. In patients infected with L. loa who have elevated levels of microfilaremia, ivermectin has been associated with the development of fatal encephalopathy (57) and so should be avoided. Ivermectin (pregnancy category C) has been shown to be teratogenic in mice, rats, and rabbits when given in repeated doses of 0.2, 8.1, and 4.5 times the maximum recommended human dose, respectively. Teratogenicity was characterized in the three species by cleft palate; clubbed forepaws were additionally observed in rabbits. These developmental effects were found only at or near doses that were maternotoxic to the pregnant female (14, 44). There are insufficient data to recommend its use in pregnant women, although the risk of fetal damage in 203 pregnant women inadvertently treated with ivermectin was no greater than that in controls, and it has been suggested that ivermectin can be given safely after the first trimester (58–60). It is excreted in breast milk in low concentrations, so it should be avoided in lactating women when possible. It is not recommended in children weighing 50% is excreted unchanged in the urine. There is negligible protein binding, and it is widely distrib-

149. Antiparasitic Agents n TABLE 7

2535

Major indications for DECa

Indication

Usual dose

Reported efficacy

Loa loa

Up to 9 mg/kg/day in 3 doses for 12 days

Wuchereria bancrofti and Brugia spp. infections

Up to 6 mg/kg/day in 3 doses for 12 days (or repeated single doses)

Few large trials, but single course is curative in 90–99% reduction in microfilaremia, but often need additional courses to eradicate adult worms

a

Adapted from reference 1.

uted in tissues. It readily crosses the blood-brain barrier. Peak levels in plasma of 100 to 150 μg/liter at 1 to 2 h have been reported after a single 0.5-mg/kg dose (62). The halflife of DEC is 2 to 10 h. Renal excretion is reduced in the presence of an alkaline urinary pH, and dose reductions are required in patients with renal impairment. Spectrum of activity. DEC is an effective microfilaricidal drug against W. bancrofti, B. malayi, Brugia timori, Onchocerca volvulus, L. loa, and M. streptocerca, but it has little or no effect on Mansonella perstans or M. ozzardi microfilariae (4). It has been the drug of choice for lymphatic filariasis for the last 50 years. It is used both for individual therapy for filarial infections and in mass community chemotherapy programs, either alone or in combination with ivermectin, albendazole, or doxycycline. It is predominantly a microfilaricidal agent, although it has some macrofilaricidal activity in W. bancrofti, B. malayi, B. timori, and L. loa infections (63–65). DEC has also been used in the treatment of toxocaral visceral larva migrans, but albendazole is now the preferred agent because of its better safety profile. Adverse effects. The side effects of DEC include mild headache, dizziness, anorexia, nausea, and arthralgias. Administration of the drug to individuals with filarial infections can induce adverse results that are not due to the drug itself but instead are related to host responses, in part to release of Wolbachia endosymbionts from filariae, following damage to adult worms (local reactions) and death of microfilariae (systemic reactions). These reactions tend to be relatively mild with lymphatic filariasis and infrequent with loiasis but can be severe with onchocerciasis or with heavy L. loa infections and result in intense pruritis, rash, fever, hypotension, and encephalopathy. The potentially fatal Mazzotti reaction and serious ophthalmic adverse effects can occur following treatment of O. volvulus with DEC, so DEC is contraindicated for onchocercal infections. For heavy L. loa infections, DEC is contraindicated to avoid potentially fatal encephalopathy. Animal studies have not shown DEC to be teratogenic, but it may increase the risk of abortion and so should be avoided in pregnancy when possible (14, 66). It is not excreted in breast milk and is safe during lactation.

worms become dislodged from the intestinal wall and are expelled in the feces by normal peristalsis. Pharmacokinetics. Pyrantel is administered orally but is almost insoluble in water and is therefore poorly absorbed from the gastrointestinal tract. Peak levels in serum occur after 1 to 3 h. More than 50% is excreted unchanged in the feces. The absorbed drug is partially metabolized in the liver. There is no significant interaction with food. Spectrum of activity. Pyrantel has excellent efficacy in the treatment of ascariasis and hookworm and pinworm infections. It also has some activity against Trichostrongylus. It is not active against T. trichiura. Adverse effects. Although pyrantel is generally well tolerated, it can lead to adverse reactions, including anorexia, nausea, vomiting, abdominal cramps, and diarrhea. It has also been associated with neurotoxic effects, including headache, dizziness, drowsiness, and insomnia. Transient increases in hepatic enzymes have also been reported, and one study reported nephrotic syndrome temporally related to its use (67). Animal studies have not shown adverse effects in the fetus, and it has been used during pregnancy in humans without harmful fetal effects (pregnancy category C) (68). It is not recommended for children 99%), and its urinary elimination half-life is 7.3 h (76). Tizoxanide then undergoes glucuronidation to form tizoxanide glucuronide. Nitazoxanide is not detected in plasma, urine, bile, or feces, but tizoxanide is found in plasma, urine, bile, and feces and tizoxanide glucuronide is found in plasma, bile, and urine (74). The pharmacokinetics of nitazoxanide in patients with impaired liver or renal function has not been studied, and it must be administered with caution to these patients (69). Spectrum of activity. While nitazoxanide has been reported to have activity against a broad range of parasites, it is FDA approved only for treatment of Cryptosporidium species and G. duodenalis, and its role in treating other infections is ill defined. Adverse effects. Nitazoxanide is well tolerated, with adverse effects similar to those of placebo. Mild and transient side effects have been seen in only 3 to 4% of patients, principally related to the gastrointestinal tract (abdominal pain, diarrhea, and nausea) (77). No significant adverse effects on electrocardiography, hematology, clinical chemistry, or urinalysis parameters in humans have been noted (71, 78). There are minimal data on the safety of nitazoxanide in pregnant (pregnancy category B) or lactating women (69).

ANTIMALARIALS A number of agents are approved for use for treatment of malaria. The most commonly recommended agents are shown in Table 9.

Quinoline Derivatives The quinoline derivatives can be divided into four groups: the 4-aminoquinolines; the cinchona alkaloids; synthetic compounds, such as mefloquine and halofantrine; and the

8-aminoquinolines. A related drug, piperaquine, which is a bisquinoline, is now often used in combination with artemisinin derivatives for treatment of malaria. Although it is used in Europe, it is not approved by the U.S. FDA.

4-Aminoquinolines The 4-aminoquinolines include chloroquine, hydroxychloroquine, and amodiaquine. Chloroquine is the most widely used of these agents. It is an inexpensive, safe drug that has been used extensively for treatment and prophylaxis of all Plasmodium species that infect humans, although resistance to chloroquine in Plasmodium falciparum is prevalent globally in most malarious regions. Hydroxychloroquine is a related synthetic compound with an identical clinical spectrum, similar pharmacokinetics, and similar adverse effects. Amodiaquine is another related agent with the same mechanism of action and spectrum of activity. It is reported to be more effective than chloroquine for parasite clearance, but its use has been restricted due to uncommon serious side effects, as noted below. It has not been approved by the U.S. FDA (1). Mechanism of action. The main mechanism of action of the 4-aminoquinolines is thought to be via nonenzymatic inhibition of heme polymerization. Asexual intraerythrocytic malaria parasites actively concentrate quinoline ring compounds within hemoglobin-containing vesicles. In the absence of drug, plasmodia degrade host erythrocyte hemoglobin to provide amino acid nutrients essential for parasite growth. The degradation of hemoglobin produces free heme, which is stored as ferriprotoporphyrin IX within the red blood cell. Ferriprotoporphyrin IX is toxic to the parasite and is usually polymerized into nontoxic malaria pigment (hemozoin). In the presence of drug, there is inhibition of the conversion of heme into hemozoin, leading to the accumulation of products toxic to the parasite and resulting in parasite death (79). These agents also inhibit protein synthesis by inhibiting incorporation of phosphate into DNA and RNA and by inhibiting DNA and RNA polymerases (13). Chloroquine also raises the pH of the vesicle (80).

TABLE 9 Treatment regimens for malaria Organism Plasmodium vivax and Plasmodium ovale Plasmodium falciparum Plasmodium malariae Plasmodium knowlesi

Primary agent used for treatment Artemisinin derivative or Malarone or chloroquine, followed by primaquine Artemisinin derivative or Malarone Chloroquine Artemisinin derivative or Malarone or chloroquine

Alternative agents Quinine sulfate plus doxycycline or plus clindamycin

149. Antiparasitic Agents n

Pharmacokinetics. The 4-aminoquinolines are extensively distributed in tissues and are characterized by a long elimination half-life. Despite similarities in their chemical structures, these drugs show differences in their biotransformation and routes of elimination (81). Chloroquine is available in oral and parenteral forms. Many different formulations are manufactured worldwide. Chloroquine is rapidly absorbed from the gastrointestinal tract after oral administration and has oral bioavailability exceeding 75%. Food has variable effects on absorption. The drug is distributed extensively in body tissues and reaches high levels within the brain (82). Chloroquine binds to melanin-containing cells in the skin and eye, so it can also reach high levels at these sites. There is marked variability in peak concentrations in plasma between individuals, but within 3 h of initiating standard oral treatment doses (10 mg chloroquine base/kg, followed by three doses of 5 mg/ kg at 6, 24, and 48 h), concentrations in blood remain above 1 mmol/liter for at least 4 days (83). It is ∼60% protein bound and has a half-life of 3 to 6 days. Approximately 30 to 50% of the drug is metabolized to inactive compounds in the liver, and the remainder is excreted in the urine. Treatment reduction (usually 50% of the normal dose) is required in patients with severe renal or hepatic failure. It is not dialyzable. In contrast, amodiaquine is a prodrug and is almost entirely metabolized to a biologically active metabolite, desethylamodiaquine, following oral administration. Otherwise, it has pharmacokinetic properties similar to those of chloroquine but has a smaller volume of distribution. Spectrum of activity. The 4-aminoquinolines are efficient and rapidly acting blood schizonticides. They can be used in both the treatment and prophylaxis of infection with susceptible strains of all Plasmodium species. They have no effect on tissue schizonts or exoerythrocytic stages. They are gametocytocidal for Plasmodium vivax and Plasmodium malariae but have minimal effect on P. falciparum gametocytes. Following infections with P. vivax or Plasmodium ovale, primaquine is also needed to eradicate liver hypnozoites and prevent relapses of infection. Chloroquine currently is still a first-line option for therapy for some P. vivax, P. malariae, and P. ovale infections, but increasing resistance among P. vivax isolates globally, and especially among P. vivax infections acquired in Indonesia, East Timor, Papua New Guinea, and the Solomon Islands, is emerging. It also remains the treatment of choice for susceptible P. falciparum strains, although P. falciparum strains from almost all areas of the world have developed resistance to it and so it is now rarely used for P. falciparum. It is also effective against Plasmodium knowlesi infections (84). Because of its potential toxicity, amodiaquine is not recommended for prophylaxis of malaria and is generally not used as first-line treatment. However, it results in faster parasite clearance and more rapid resolution of symptoms than those with chloroquine, and it may be effective in some cases of chloroquine-resistant malaria, so it is used as an alternative treatment regimen in some areas. Chloroquine is also active against E. histolytica trophozoites but is rarely used for this indication, as the nitro-5imidazoles are the drugs of choice. Adverse effects. Chloroquine has a bitter taste. It is generally well tolerated at the doses required for malaria prevention or treatment, even when taken for prolonged periods. However, it can lead to nausea, abdominal discomfort, dizziness, retinal pigmentation, blurred vision, electro-

2537

cardiographic changes, muscular weakness, and, rarely, transient psychiatric symptoms. It can also cause severe pruritis, particularly in African blacks. Irreversible neuroretinitis can result if it is taken at high doses for prolonged periods. If taken as an overdose, it can cause shock, arrhythmia, and death. At the doses used for malaria treatment or prophylaxis, chloroquine has rarely been reported to cause adverse congenital effects (pregnancy category C) (85). However, affinity for melanin-containing tissues, such as the retina, iris, and choroid of the eye, has been reported, and definitive delineation of fetal risk remains undefined. It is used commonly for treatment and prophylaxis of malaria in pregnant women, without evidence of teratogenicity, and it is generally agreed that the benefits of preventing and treating malaria in pregnant women outweigh the potential fetal risks. Chloroquine is excreted in small amounts in breast milk. Amodiaquine is more palatable than chloroquine and seems to cause less itching. However, serious adverse events, including agranulocytosis, aplastic anemia, and drug-induced hepatitis, have been reported. These have occurred predominantly following long-term amodiaquine use (mean, 7 to 8 weeks) for malaria prophylaxis. While short-term treatment regimens are thought to be safe (86), this drug is now used uncommonly.

Cinchona Alkaloids The cinchona alkaloids, quinine and quinidine, contain a quinoline ring. Quinidine is the diastereoisomer of quinine. Quinine was originally extracted from the bark of the South American cinchona tree, but a synthetic form is now available, usually as a quinine sulfate salt. Quinidine is more active than quinine, but it is also more cardiotoxic. Mechanism of action. The exact target of cinchona alkaloids is unknown. They are thought to act by forming complexes with ferriprotoporphyrin IX, thereby interfering with hemoglobin digestion and resulting in cell lysis and death of schizonts (87). They also interfere with the function of plasmodial DNA and inhibit the synthesis of parasite nucleic acids and proteins. Quinine also interacts with certain fatty acids present in parasitized erythrocytes, preventing red blood cell lysis and interrupting schizont maturation (13). Additionally, it increases intracellular pH, resulting in lethal effects on the parasite. Pharmacokinetics. Quinine is available for oral administration as a sulfate salt and for parenteral administration as quinine dihydrochloride. It is >80% absorbed from the gastrointestinal tract following oral doses. It is widely distributed in body tissues, but concentrations in CSF are 90% protein bound. Quinine is metabolized in the liver, and the native drug and its metabolites are excreted in the urine (82). It has a short half-life of 8 to 12 h, necessitating multiple daily doses. After a single dose of 650 mg of quinine sulfate, peak concentrations in serum are ∼3.2 mg/liter in healthy individuals but are higher (8.4 mg/liter) in patients with malaria. Intravenous quinine is used in many countries when oral therapy cannot be tolerated, but quinidine gluconate is considered the parenteral drug of choice in the United States (1). Both agents have similar pharmacokinetic properties. The pharmacokinetic properties of the cinchona alkaloids are considerably altered in patients with malaria, with a reduction in clearance that is proportional to the severity

2538 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

of disease. Consequently, doses should be decreased by 30 to 50% after the third day of treatment to avoid accumulation of drug in seriously ill patients (88). Drug levels may also be increased by administration with foods that alkalinize the urine, because increased tubular reabsorption results. Caution is recommended for patients with significant liver impairment, and dose reduction is required if there is renal impairment. Both agents are partially dialyzable. Spectrum of activity. The cinchona alkaloids can be used in the treatment of all Plasmodium species that infect humans. Their main indication is for chloroquine-resistant P. falciparum. Oral therapy is indicated for uncomplicated malaria, but intravenous formulations of quinine dihydrochloride or quinidine gluconate are used in severe infections. Quinine and quinidine are blood schizonticides but have little effect on sporozoites or preerythrocytic forms of the parasite. Consequently, they do not eradicate P. vivax or P. ovale hypnozoites in the liver. They also are not gametocytocidal against P. falciparum. Although resistance to these agents has emerged in Southeast Asia and Africa, they remain useful drugs for the treatment of malaria worldwide; however, over recent years they have been replaced by artemisinin derivatives for first-line malaria treatment. Quinine is also used for the treatment of babesiosis. It is ineffective when used as a single agent but can be given together with clindamycin or azithromycin (1). Adverse effects. Quinine has an extremely bitter taste and can be associated with nausea, vomiting, and epigastric pain. It also often leads to the symptom complex of cinchonism (nausea, tinnitus, dysphoria, and reversible high-tone deafness). Quinine can also cause hyperinsulinemic hypoglycemia, especially in children and pregnant women with severe malaria, as it increases release of insulin from the pancreas. It has also been associated with massive hemolysis in patients with heavy P. falciparum infections. Agranulocytosis, thrombocytopenia, retinopathy, and tongue discoloration are additional adverse effects that have been described. Overdose of quinine can lead to ataxia, convulsions, and coma. When used as treatment for severe malaria, intravenous quinidine is associated with cardiac arrhythmias. It prolongs the QT interval, widens the QRS complex, and prolongs the PR interval. It can therefore lead to hypotension and ventricular arrhythmias, including torsades de pointes. Consequently, it should be administered only in an intensive care setting with cardiac monitoring. As with quinine, administration can also result in blood dyscrasias and cinchonism. Despite reports of congenital defects following administration of quinine during pregnancy, it can be administered during pregnancy when the benefits of maternal treatment outweigh the potential fetal risks (pregnancy category C) (89, 90). Quinidine has not been reported to be teratogenic. Quinine can have an abortificient effect and can lead to induction of labor. It is excreted in small amounts in breast milk but can be administered during breast feeding when necessary.

Synthetic Quinoline Compounds Mefloquine Mefloquine is a synthetic 4-quinoline methanol compound structurally related to quinine.

the parasitized erythrocyte (13). Its action is thought to rely on interference with the digestion of hemoglobin during the blood stages of the malaria life cycle, likely via a mechanism similar to that of quinine (87). It does not inhibit protein synthesis. Pharmacokinetics. Mefloquine is available for oral administration only. Food enhances bioavailability, and it should not be taken on an empty stomach. It is >85% absorbed following oral administration and is concentrated within red blood cells (91). It is >95% protein bound and has a half-life of 2 to 4 weeks. Because of its long half-life, mefloquine is frequently used for prophylaxis of malaria, as a once-weekly dose. However, when mefloquine is administered weekly, it requires about 8 weeks before steady-state drug levels are reached, so a loading dose is often recommended. Peak concentrations in plasma occur at 6 to 24 h, and following a single dose of 500 mg or 1,000 mg orally, they are 430 and 800 μg/liter, respectively. In healthy volunteers, a dose of 250 mg once weekly produces maximum steady-state plasma concentrations of 1,000 to 2,000 μg/ liter, which are reached after 7 to 10 weeks. It is highly lipophilic, is widely distributed throughout the body, and can cross the blood-brain barrier. Mefloquine is metabolized in the liver and excreted through the bile and feces. There are no specific recommendations regarding the need for dosage adjustment in patients with renal or hepatic failure. It is not dialyzable. Spectrum of activity. Mefloquine is active against the erythrocytic schizonts of all Plasmodium species causing human malaria, and it has been used for both chemoprophylaxis and therapy. Since weekly administration is sufficient for chemoprophylaxis, it is convenient for use in travelers to areas where malaria is endemic. Its main utility in malaria treatment results from its activity against most chloroquineresistant P. falciparum strains, although resistance has been recognized, particularly in some areas of Southeast Asia. When used for therapy, it should be combined with another agent, usually an artemisinin derivative. Mefloquine does not kill tissue schizonts, so patients infected with P. vivax should subsequently be treated with an 8-aminoquinoline. It also has no effect on gametocytes. Adverse effects. Adverse reactions to mefloquine include nausea and vomiting, agranulocytosis, and aplastic anemia, as well as central nervous system (CNS) effects such as dysphoria, dizziness, disturbed sleep, nightmares, and ataxia. Severe neuropsychiatric reactions, including delirium and seizures, have been reported occasionally and are thought to occur in approximately 1:200 to 1:1,300 patients treated for acute falciparum malaria (92, 93). Mefloquine can also potentiate dysrhythmias in individuals on β-blockers. Mefloquine is teratogenic in high doses in animals, but reports on humans do not support teratogenic effects (pregnancy category B) (94). A possible higher rate of spontaneous abortion has been suggested, so it is generally avoided in the first trimester of pregnancy if possible. However, limited data suggest that it is probably safe to use even during the first trimester, and it can be used in later stages of pregnancy if the benefits outweigh the potential risks (95). It is excreted in low concentrations in breast milk but can be used in lactating women when necessary.

Halofantrine Mechanism of action. Mefloquine interacts both with host cell phospholipids and with ferriprotoporphyrin IX of

Halofantrine is a synthetic phenanthrene-methanol compound. It has not been approved by the U.S. FDA (1),

149. Antiparasitic Agents n

and because of its potential cardiac side effects, it has limited indications for use. Mechanism of action. Halofantrine has activity against the asexual erythrocytic stages of malaria parasites, although the exact mechanism of action is unclear. Pharmacokinetics. Halofantrine is available only for oral administration but has variable bioavailability. Absorption is enhanced by administration with fatty food, but because high levels in blood enhance toxicity, it is recommended for administration on an empty stomach. After three doses of 500 mg of halofantrine hydrochloride (at 0, 6, and 12 h), a maximum concentration in plasma of 896 μg/liter was reported (96). Halofantrine is metabolized in the liver to an active metabolite, N-desbutylhalofantrine. The half-lives of halofantrine and its metabolite are 6 to 10 days and 3 to 4 days, respectively. It is excreted mainly in the feces. Spectrum of activity. Halofantrine is efficacious in the treatment of P. vivax and P. falciparum malaria, but data concerning P. ovale and P. malariae are limited (97). It is not recommended for prophylaxis of malaria because of toxicity. It is active against blood-stage schizonts only and appears to have no effect against sporozoites, gametocytes, or tissue-stage parasites. Halofantrine is more active than mefloquine, but cross-resistance between these drugs occur. Its expense and potential toxicity also limit its use. Adverse effects. Halofantrine leads to gastrointestinal adverse effects, including nausea, vomiting, diarrhea, and abdominal pain. It also has potential cardiovascular toxicity and causes concentration-dependent prolongation of the QT interval. It is therefore contraindicated in patients with long QT syndrome, as it can lead to cardiac arrest (98). It can also lead to pruritis and hepatic enzyme elevations. Halofantrine is contraindicated in pregnancy. The degree of excretion in breast milk is unknown, and it is not advised for use in lactating women.

Lumefantrine Lumefantrine is a drug with a similar structure to that of halofantrine. It is widely used as a long-acting partner drug to artemether in a fixed-dose combination for malaria. Mechanism of action. The exact mechanism by which lumefantrine exerts its antimalarial effect is unknown. However, it seems to inhibit the formation of β-hematin by forming a complex with hemin and inhibits nucleic acid and protein synthesis. Pharmacokinetics. The oral bioavailability of lumefantrine is highly variable and increases up to 3- to 4-fold when it is taken with a high-fat meal. Peak levels in plasma are seen after 6 to 8 h. Lumefantrine is 99.7% protein bound, and its half-life is 3 to 6 days. It is highly lipophilic and has an apparent large volume of distribution. Peak levels in plasma vary considerably, but after administration of six tablets containing 2,780 mg of lumefantrine, median levels in plasma of 8 to 9 μg/ml were reported (99). Lumefantrine is extensively metabolized in the liver, and the major metabolite found in plasma is desbutyl-lumefantrine. It should be used with caution in patients with severe renal or hepatic failure. Spectrum of activity. In its fixed-dose combination with artemether, lumefantrine has efficacy against all Plasmodium species.

2539

Adverse effects. Lumefantrine is well tolerated, with rare mild adverse reactions such as diarrhea, nausea, abdominal pain, and vomiting. There is no evidence of significant cardiotoxicity associated with lumefantrine use, but it is recommended that it be avoided in patients at risk for QT prolongation. Artemether-lumefantrine has been assigned to pregnancy category C by the FDA. There are no human data on the excretion of lumefantrine into breast milk, but animal data suggest some excretion. The effects in the nursing infant are unknown, and it should be used with caution in lactating women.

8-Aminoquinolines The 8-aminoquinolines are primaquine and tafenoquine (WR 238,605). Tafenoquine is not yet commercially available and has not been approved by the U.S. FDA (1). Mechanism of action. The 8-aminoquinolines interfere with parasite mitochondrial enzymes involved in energy production. They also have an inhibitory action against DNA, although the exact mechanism by which this occurs in unclear (13). An active metabolite of the drug is thought to interrupt the mitochondrial transport system and pyrimidine synthesis in hypnozoites (100). Pharmacokinetics. The 8-aminoquinolines are well absorbed after oral administration, with >90% bioavailability. Primaquine is rapidly metabolized in the liver, and its halflife is 4 to 9 h, so it needs to be administered daily. Following a single 45-mg oral dose, a mean peak level in serum of 153.3 μg/liter was observed after 2 to 3 h (101). It is found at relatively low concentrations in most body sites. Tafenoquine has a longer half-life of 2 to 4 weeks. Weekly or possibly monthly doses seem to be sufficient for prophylaxis, thus making it better tolerated than primaquine (102). A single dose of tafenoquine may be sufficient for prevention of relapse following P. vivax infections (103). Optimal dosefinding studies are being performed. Spectrum of activity. Primaquine is less active against blood-stage malarial forms than most other antimalarial agents are, but it is very active against preerythrocytic sporozoites and exoerythrocytic tissue schizonts of all malarial species. Its main use is to prevent relapse of P. vivax and P. ovale infections from latent hypnozoites following treatment with chloroquine. Additionally, it is gametocytocidal against Plasmodium, especially P. falciparum, and can interrupt transmission of malaria. It is also an effective causal prophylactic agent but traditionally has been used infrequently in this way for travelers. Tafenoquine is reported to be more active than primaquine and has higher schizonticidal activity (104). Adverse effects. Mild gastrointestinal side effects, including nausea and abdominal pain, are common following 8-aminoquinoline administration. They should not be used in people with glucose-6-phosphate dehydrogenase deficiency, as they can induce hemolysis. Patients with NADH methemoglobin reductase deficiency are at risk of developing methemoglobinemia. Primaquine also occasionally causes arrhythmias. Interference with visual accommodation has also been reported. These agents should also not be used during pregnancy or lactation because of the potential risk of hemolytic effects in the fetus.

Artemisinin (Qinghasou) Derivatives Artemisinin is an extract from the Chinese herbal plant Artemisia annua, also known as qinghasou. It is a sesquiterpene

2540 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

lactone peroxide. Synthetic derivatives include artemether, dihydroartemisinin, arteether, and artesunate. Although not officially approved for use by the U.S. FDA, the intravenous formulation of artesunate is available via the CDC under an IND for patients with severe malaria. Mechanism of action. Artemisinin and its derivatives act mainly against the asexual erythrocytic stages of malaria parasites. They have an antiparasitic effect, particularly on young, ring-form parasites, leading to their clearance and preventing development of more mature pathogenic forms. They bind to the parasite membrane and to ferriprotoporphyrin IX, so they are highly concentrated within parasites and reach 100 to 300 times higher concentrations in P. falciparum-infected red cells than in uninfected cells (105). By binding iron in the malarial pigment, they lead to the production of toxic oxidative free radicals that damage parasite organelles and alkylate parasitic proteins, leading to inhibition of protein synthesis and ultimately to parasite death. Pharmacokinetics. The derivatives of artemisinin have greater solubility than artemisinin and consequently have been developed for easier administration by a variety of routes. Artesunate is water-soluble and can be given intravenously, intramuscularly, orally, or by suppository. Artemether and arteether are oil-soluble and are available in tablet, capsule, and intramuscular injection forms. Although artesunate is the most potent in vitro, there is no apparent clinical difference in efficacy between the formulations. The artemisinin derivatives have a short half-life of 90%. It is ∼40% protein bound. It has a half-life of ∼12 h and has good tissue penetration (163). Nifurtimox is administered orally but has poor oral bioavailability. It is metabolized in the liver and has a half-life of ∼3 h. Dose reduction is advised for patients with significant hepatic or renal impairment, but no specific guidelines exist. Spectrum of activity. Both benznidazole and nifurtimox are used for the treatment of acute T. cruzi infection (Chagas’ disease). Neither agent has demonstrated efficacy in late stages of disease, and indications for treatment of chronic infection remain controversial but are expanding (164– 166). No randomized trial has evaluated the comparative safety and efficacy of nifurtimox and benznidazole in adults (167). Nifurtimox is now also increasingly being used in combination with eflornithine for first-line treatment of T. b. gambiense infection. It also has been shown to have some activity in leishmaniasis but is not routinely used for this indication.

2545

Adverse effects. Side effects are common with benznidazole and are seen in up to 40% of treated individuals, commonly including vomiting, abdominal pain, peripheral neuropathy, rash, and pruritis. Bone marrow suppression and neuropsychiatric reactions have also been reported. Nifurtimox has significant side effects that preclude the completion of therapy in many patients. Adverse effects include anorexia, nausea, rash, headache, sleep disturbance, peripheral neuropathy, and myalgias. Less-frequent but more severe toxicities include psychosis and convulsions. Benznidazole crosses the placenta, but there are minimal data regarding teratogenic effects for either agent in either animals or humans (168). Nifurtimox is detected in breast milk, so caution is recommended during breast feeding (169). Similarly, safety data for benznidazole and lactation are lacking, so withholding treatment during breast feeding is again recommended.

REFERENCES 1. The Medical Letter. 2013. Drugs for Parasitic Infections. The Medical Letter, Inc, New Rochelle, NY. 2. Mackenzie CD, Geary TG. 2011. Flubendazole: a candidate macrofilaricide for lymphatic filariasis and onchocerciasis field programs. Expert Rev Anti Infect Ther 9:497–501. 3. Ceballos L, Virkel G, Elissondo C, Canton C, Canevari J, Murno G, Denegri G, Lanusse C, Alvarez L. 2013. A pharmacology-based comparison of the activity of albendazole and flubendazole against Echinococcus granulosus metacestode in sheep. Acta Trop 127:216–225. 4. van den Enden E. 2009. Pharmacotherapy of helminth infection. Expert Opin Pharmacother 10:435–451. 5. Calvopina M, Guderian RH, Paredes W, Chico M, Cooper PJ. 1998. Treatment of human pulmonary paragonimiasis with triclabendazole: clinical tolerance and drug efficacy. Trans R Soc Trop Med Hyg 92:566–569. 6. Villegas F, Angles R, Barrientos R, Barrios G, Valero MA, Hamed K, Grueninger H, Ault SK, Montresor A, Engels D, Mas-Coma S, Gabrielli AF. 2012. Administration of triclabendazole is safe and effective in controlling fascioliasis in an endemic community of the Bolivian Altiplano. PLoS Negl Trop Dis 6:e1720. doi:10.1371/journal.pntd.0001720. 7. Cabada MM, White AC, Jr. 2012. New developments in epidemiology, diagnosis, and treatment of fascioliasis. Curr Opin Infect Dis 25:518–522. 8. Winkelhagen AJ, Mank T, de Vries PJ, Soetekouw R. 2012. Apparent triclabendazole-resistant human Fasciola hepatica infection, the Netherlands. Emerg Infect Dis 18:1028–1029. 9. Fairweather I. 2009. Triclabendazole progress report, 2005–2009: an advancement of learning? J Helminthol 83:139–150. 10. Keiser J, Engels D, Buscher G, Utzinger J. 2005. Triclabendazole for the treatment of fascioliasis and paragonimiasis. Expert Opin Investig Drugs 14:1513–1526. 11. Lacey E. 1990. Mode of action of benzimidazoles. Parasitol Today 6:112–115. 12. Gilman A, Rall TW, Nies AS (ed). 1990. Goodman and Gilman’s The Pharmacological Basis of Therapeutics, 8th ed. Pergamon Press, New York, NY. 13. Frayha GJ, Smyth JD, Gobert JG, Savel J. 1997. The mechanisms of action of antiprotozoal and anthelmintic drugs in man. Gen Pharmacol 28:273–299. 14. de Silva N, Guyatt H, Bundy D. 1997. Anthelmintics. A comparative review of their clinical pharmacology. Drugs 53:769–788. 15. Edwards G, Breckenridge AM. 1988. Clinical pharmacokinetics of anthelmintic drugs. Clin Pharmacokinet 15:67–93. 16. Marriner SE, Morris DL, Dickson B, Bogan JA. 1986. Pharmacokinetics of albendazole in man. Eur J Clin Pharmacol 30:705–708.

2546 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

17. Bogan JA, Marriner S. 1980. Analysis of benzimidazoles in body fluids by high-performance liquid chromatography. J Pharm Sci 69:422–423. 18. Jung H, Hurtado M, Sanchez M, Medina MT, Sotelo J. 1992. Clinical pharmacokinetics of albendazole in patients with brain cysticercosis. J Clin Pharmacol 32:28–31. 19. Jung H, Hurtado M, Medina MT, Sanchez M, Sotelo J. 1990. Dexamethasone increases plasma levels of albendazole. J Neurol 237:279–280. 20. Garcia HH, Lescano AG, Lanchote VL, Pretell EJ, Gonzales I, Bustos JA, Takayanagui OM, Bonato PS, Horton J, Saavedra H, Gonzalez AE, Gilman RH, Cysticercosis Working Group in Peru. 2011. Pharmacokinetics of combined treatment with praziquantel and albendazole in neurocysticercosis. Br J Clin Pharmacol 72:77–84. 21. Lima RM, Ferreira MA, de Jesus Ponte Carvalho TM, Dumet Fernandes BJ, Takayanagui OM, Garcia HH, Coelho EB, Lanchote VL. 2011. Albendazole-praziquantel interaction in healthy volunteers: kinetic disposition, metabolism and enantioselectivity. Br J Clin Pharmacol 71: 528–535. 22. Morris DL, Chinnery JB, Georgiou G, Stamatakis G, Golematis B. 1987. Penetration of albendazole sulphoxide into hydatid cysts. Gut 28:75–80. 23. Moskopp D, Lotterer E. 1993. Concentrations of albendazole in serum, cerebrospinal fluid and hydatidous brain cyst. Neurosurg Rev 16:35–37. 24. Keiser J, Utzinger J. 2008. Efficacy of current drugs against soil-transmitted helminth infections: systematic review and meta-analysis. JAMA 299:1937–1948. 25. Opatrny L, Prichard R, Snell L, Maclean JD. 2005. Death related to albendazole-induced pancytopenia: case report and review. Am J Trop Med Hyg 72:291–294. 26. Lillie P, McGann H. 2010. Empiric albendazole therapy and new onset seizures—a cautionary note. J Infect 60:403– 404. (Author’s reply, 60:404–405.) 27. Ramos-Zuniga R, Perez-Gomez HR, Jauregui-Huerta F, del Sol Lopez-Hernandez M, Valera-Lizarraga JE, PazVelez G, Becerra-Valdivia A. 2013. Incidental consequences of antihelmintic treatment in the central nervous system. World Neurosurg 79:149–153. 28. Auer H, Kollaritsch H, Juptner J, Aspock H. 1994. Albendazole and pregnancy. Appl Parasitol 35:146–147. 29. Abdel-tawab AM, Bradley M, Ghazaly EA, Horton J, elSetouhy M. 2009. Albendazole and its metabolites in the breast milk of lactating women following a single oral dose of albendazole. Br J Clin Pharmacol 68:737–742. 30. Brindley PJ, Sher A. 1990. Immunological involvement in the efficacy of praziquantel. Exp Parasitol 71:245–248. 31. Doenhoff MJ, Cioli D, Utzinger J. 2008. Praziquantel: mechanisms of action, resistance and new derivatives for schistosomiasis. Curr Opin Infect Dis 21:659–667. 32. Sotelo J, Jung H. 1998. Pharmacokinetic optimisation of the treatment of neurocysticercosis. Clin Pharmacokinet 34:503–515. 33. Kaojarern S, Nathakarnkikool S, Suvanakoot TU. 1989. Comparative bioavailability of praziquantel tablets. DICP 23:29–32. 34. Mohamed AE, Yasawy MI, Al Karawi MA. 1998. Combined albendazole and praziquantel versus albendazole alone in the treatment of hydatid disease. Hepatogastroenterology 45:1690–1694. 35. Cobo F, Yarnoz C, Sesma B, Fraile P, Aizcorbe M, Trujillo R, Diaz-de-Liano A, Ciga MA. 1998. Albendazole plus praziquantel versus albendazole alone as a pre-operative treatment in intra-abdominal hydatisosis caused by Echinococcus granulosus. Trop Med Int Health 3:462–466. 36. Bygott JM, Chiodini PL. 2009. Praziquantel: neglected drug? Ineffective treatment? Or therapeutic choice in cystic hydatid disease? Acta Trop 111:95–101.

37. Lee JM, Lim HS, Hong ST. 2011. Hypersensitive reaction to praziquantel in a clonorchiasis patient. Korean J Parasitol 49:273–275. 38. Kyung SY, Cho YK, Kim YJ, Park JW, Jeong SH, Lee JI, Sung YM, Lee SP. 2011. A paragonimiasis patient with allergic reaction to praziquantel and resistance to triclabendazole: successful treatment after desensitization to praziquantel. Korean J Parasitol 49:73–77. 39. Hewagama SS, Darby JD, Sheorey H, Daffy JR. 2010. Seizures related to praziquantel therapy in neurocysticercosis. Med J Aust 193:246–247. 40. Olds GR. 2003. Administration of praziquantel to pregnant and lactating women. Acta Trop 86:185–195. 41. Adam I, Elwasila el T, Homeida M. 2004. Is praziquantel therapy safe during pregnancy? Trans R Soc Trop Med Hyg 98:540–543. 42. Allen HE, Crompton DW, de Silva N, LoVerde PT, Olds GR. 2002. New policies for using anthelmintics in high risk groups. Trends Parasitol 18:381–382. 43. Sutherland IH, Campbell WC. 1990. Development, pharmacokinetics and mode of action of ivermectin. Acta Leiden 59:161–168. 44. Campbell WC. 1993. Ivermectin, an antiparasitic agent. Med Res Rev 13:61–79. 45. Fox LM. 2006. Ivermectin: uses and impact 20 years on. Curr Opin Infect Dis 19:588–593. 46. Geary TG, Moreno Y. 2012. Macrocyclic lactone anthelmintics: spectrum of activity and mechanism of action. Curr Pharm Biotechnol 13:866–872. 47. Ottesen EA, Campbell WC. 1994. Ivermectin in human medicine. J Antimicrob Chemother 34:195–203. 48. Fusco DN, Downs JA, Satlin MJ, Pahuja M, Ramos L, Barie PS, Fleckenstein L, Murray HW. 2010. Non-oral treatment with ivermectin for disseminated strongyloidiasis. Am J Trop Med Hyg 83:879–883. 49. Grein JD, Mathisen GE, Donovan S, Fleckenstein L. 2010. Serum ivermectin levels after enteral and subcutaneous administration for Strongyloides hyperinfection: a case report. Scand J Infect Dis 42:234–236. 50. Baraka OZ, Mahmoud BM, Marschke CK, Geary TG, Homeida MM, Williams JF. 1996. Ivermectin distribution in the plasma and tissues of patients infected with Onchocerca volvulus. Eur J Clin Pharmacol 50:407–410. 51. Naquira C, Jimenez G, Guerra JG, Bernal R, Nalin DR, Neu D, Aziz M. 1989. Ivermectin for human strongyloidiasis and other intestinal helminths. Am J Trop Med Hyg 40:304–309. 52. Wen LY, Yan XL, Sun FH, Fang YY, Yang MJ, Lou LJ. 2008. A randomized, double-blind, multicenter clinical trial on the efficacy of ivermectin against intestinal nematode infections in China. Acta Trop 106:190–194. 53. Knopp S, Mohammed KA, Speich B, Hattendorf J, Khamis IS, Khamis AN, Stothard JR, Rollinson D, Marti H, Utzinger J. 2010. Albendazole and mebendazole administered alone or in combination with ivermectin against Trichuris trichiura: a randomized controlled trial. Clin Infect Dis 51:1420–1428. 54. Kraivichian K, Nuchprayoon S, Sitichalernchai P, Chaicumpa W, Yentakam S. 2004. Treatment of cutaneous gnathostomiasis with ivermectin. Am J Trop Med Hyg 71:623–628. 55. Gonzalez P, Gonzalez FA, Ueno K. 2012. Ivermectin in human medicine, an overview of the current status of its clinical applications. Curr Pharm Biotechnol 13:1103–1109. 56. Njoo FL, Beek WM, Keukens HJ, van Wilgenburg H, Oosting J, Stilma JS, Kijlstra A. 1995. Ivermectin detection in serum of onchocerciasis patients: relationship to adverse reactions. Am J Trop Med Hyg 52:94–97. 57. Twum-Danso NA. 2003. Loa loa encephalopathy temporally related to ivermectin administration reported from on-

149. Antiparasitic Agents n

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

68. 69. 70. 71.

72.

73.

74.

75.

76.

chocerciasis mass treatment programs from 1989 to 2001: implications for the future. Filaria J 2(Suppl 1):S7. doi:10.1186/1475-2883-2-S1-S7. Pacque M, Munoz B, Poetschke G, Foose J, Greene BM, Taylor HR. 1990. Pregnancy outcome after inadvertent ivermectin treatment during community-based distribution. Lancet 336:1486–1489. Goa KL, McTavish D, Clissold SP. 1991. Ivermectin. A review of its antifilarial activity, pharmacokinetic properties and clinical efficacy in onchocerciasis. Drugs 42:640–658. Gyapong JO, Chinbuah MA, Gyapong M. 2003. Inadvertent exposure of pregnant women to ivermectin and albendazole during mass drug administration for lymphatic filariasis. Trop Med Int Health 8:1093–1101. Klion AD, Ottesen EA, Nutman TB. 1994. Effectiveness of diethylcarbamazine in treating loiasis acquired by expatriate visitors to endemic regions: long-term follow-up. J Infect Dis 169:604–610. Edwards G, Awadzi K, Breckenridge AM, Gilles HM, Orme ML, Ward SA. 1981. Diethylcarbamazine disposition in patients with onchocerciasis. Clin Pharmacol Ther 30:551–557. Noroes J, Dreyer G, Santos A, Mendes VG, Medeiros Z, Addiss D. 1997. Assessment of the efficacy of diethylcarbamazine on adult Wuchereria bancrofti in vivo. Trans R Soc Trop Med Hyg 91:78–81. Ottesen EA. 1985. Efficacy of diethylcarbamazine in eradicating infection with lymphatic-dwelling filariae in humans. Rev Infect Dis 7:341–356. Nicolas L, Plichart C, Nguyen LN, Moulia-Pelat JP. 1997. Reduction of Wuchereria bancrofti adult worm circulating antigen after annual treatments of diethylcarbamazine combined with ivermectin in French Polynesia. J Infect Dis 175:489–492. Joseph CA, Dixon PA. 1984. Possible prostaglandin-mediated effect of diethylcarbamazine on rat uterine contractility. J Pharm Pharmacol 36:281–282. Ferrara P, Bersani I, Bottaro G, Vitelli O, Liberatore P, Gatto A, del Bufalo F, Romano V, Stabile A. 2011. Massive proteinuria: a possible side effect of pyrantel pamoate? Ren Fail 33:534–536. Tietze PE, Jones JE. 1991. Parasites during pregnancy. Prim Care 18:75–99. Fox LM, Saravolatz LD. 2005. Nitazoxanide: a new thiazolide antiparasitic agent. Clin Infect Dis 40:1173–1180. Anonymous. 2003. Nitazoxanide (Alinia)—a new antiprotozoal agent. Med Lett Drugs Ther 45:29–31. Gilles HM, Hoffman PS. 2002. Treatment of intestinal parasitic infections: a review of nitazoxanide. Trends Parasitol 18:95–97. Stockis A, Allemon AM, De Bruyn S, Gengler C. 2002. Nitazoxanide pharmacokinetics and tolerability in man using single ascending oral doses. Int J Clin Pharmacol Ther 40:213–220. Broekhuysen J, Stockis A, Lins RL, De Graeve J, Rossignol JF. 2000. Nitazoxanide: pharmacokinetics and metabolism in man. Int J Clin Pharmacol Ther 38:387–394. Ortiz JJ, Ayoub A, Gargala G, Chegne NL, Favennec L. 2001. Randomized clinical study of nitazoxanide compared to metronidazole in the treatment of symptomatic giardiasis in children from Northern Peru. Aliment Pharmacol Ther 15:1409–1415. Adagu IS, Nolder D, Warhurst DC, Rossignol JF. 2002. In vitro activity of nitazoxanide and related compounds against isolates of Giardia intestinalis, Entamoeba histolytica and Trichomonas vaginalis. J Antimicrob Chemother 49:103– 111. Stockis A, Deroubaix X, Lins R, Jeanbaptiste B, Calderon P, Rossignol JF. 1996. Pharmacokinetics of nitazoxanide after single oral dose administration in 6 healthy volunteers. Int J Clin Pharmacol Ther 34:349–351.

2547

77. Diaz E, Mondragon J, Ramirez E, Bernal R. 2003. Epidemiology and control of intestinal parasites with nitazoxanide in children in Mexico. Am J Trop Med Hyg 68:384–385. 78. Olliaro P, Seiler J, Kuesel A, Horton J, Clark JN, Don R, Keiser J. 2011. Potential drug development candidates for human soil-transmitted helminthiases. PLoS Negl Trop Dis 5:e1138. doi:10.1371/journal.pntd.0001138. 79. Slater AF, Cerami A. 1992. Inhibition by chloroquine of a novel haem polymerase enzyme activity in malaria trophozoites. Nature 355:167–169. 80. Krogstad DJ, Schlesinger PH, Gluzman IY. 1989. Chloroquine and acid vesicle function. Prog Clin Biol Res 313:53–59. 81. Pussard E, Verdier F. 1994. Antimalarial 4-aminoquinolines: mode of action and pharmacokinetics. Fundam Clin Pharmacol 8:1–17. 82. Krishna S, White NJ. 1996. Pharmacokinetics of quinine, chloroquine and amodiaquine. Clinical implications. Clin Pharmacokinet 30:263–299. 83. Rombo L, Bjorkman A, Sego E, Ericsson O. 1986. Whole blood concentrations of chloroquine and desethylchloroquine during and after treatment of adult patients infected with Plasmodium vivax, P. ovale or P. malariae. Trans R Soc Trop Med Hyg 80:763–766. 84. Daneshvar C, Davis TM, Cox-Singh J, Rafa’ee MZ, Zakaria SK, Divis PC, Singh B. 2010. Clinical and parasitological response to oral chloroquine and primaquine in uncomplicated human Plasmodium knowlesi infections. Malar J 9:238. doi:10.1186/1475-2875-9-238. 85. Levy M, Buskila D, Gladman DD, Urowitz MB, Koren G. 1991. Pregnancy outcome following first trimester exposure to chloroquine. Am J Perinatol 8:174–178. 86. Olliaro P, Nevill C, LeBras J, Ringwald P, Mussano P, Garner P, Brasseur P. 1996. Systematic review of amodiaquine treatment in uncomplicated malaria. Lancet 348: 1196–1201. 87. Foley M, Tilley L. 1997. Quinoline antimalarials: mechanisms of action and resistance. Int J Parasitol 27:231–240. 88. White NJ. 1985. Clinical pharmacokinetics of antimalarial drugs. Clin Pharmacokinet 10:187–215. 89. Looareesuwan S, Phillips RE, White NJ, Kietinun S, Karbwang J, Rackow C, Turner RC, Warrell DA. 1985. Quinine and severe falciparum malaria in late pregnancy. Lancet 2:4–8. 90. Silver H. 1997. Malarial infection during pregnancy. Infect Dis Clin North Am 11:99–107. 91. Karbwang J, White NJ. 1990. Clinical pharmacokinetics of mefloquine. Clin Pharmacokinet 19:264–279. 92. Hennequin C, Bouree P, Bazin N, Bisaro F, Feline A. 1994. Severe psychiatric side effects observed during prophylaxis and treatment with mefloquine. Arch Intern Med 154:2360–2362. 93. van Riemsdijk MM, van der Klauw MM, van Heest JA, Reedeker FR, Ligthelm RJ, Herings RM, Stricker BH. 1997. Neuro-psychiatric effects of antimalarials. Eur J Clin Pharmacol 52:1–6. 94. Vanhauwere B, Maradit H, Kerr L. 1998. Post-marketing surveillance of prophylactic mefloquine (Lariam) use in pregnancy. Am J Trop Med Hyg 58:17–21. 95. Nosten F, Vincenti M, Simpson J, Yei P, Thwai KL, de Vries A, Chongsuphajaisiddhi T, White NJ. 1999. The effects of mefloquine treatment in pregnancy. Clin Infect Dis 28:808–815. 96. Veenendaal JR, Parkinson AD, Kere N, Rieckmann KH, Edstein MD. 1991. Pharmacokinetics of halofantrine and N-desbutylhalofantrine in patients with falciparum malaria following a multiple dose regimen of halofantrine. Eur J Clin Pharmacol 41:161–164. 97. Bryson HM, Goa KL. 1992. Halofantrine. A review of its antimalarial activity, pharmacokinetic properties and therapeutic potential. Drugs 43:236–258.

2548 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

98. Bouchaud O, Imbert P, Touze JE, Dodoo AN, Danis M, Legros F. 2009. Fatal cardiotoxicity related to halofantrine: a review based on a worldwide safety data base. Malar J 8:289. doi:10.1186/1475-2875-8-289. 99. Ezzet F, van Vugt M, Nosten F, Looareesuwan S, White NJ. 2000. Pharmacokinetics and pharmacodynamics of lumefantrine (benflumetol) in acute falciparum malaria. Antimicrob Agents Chemother 44:697–704. 100. Warhurst DC. 1984. Why are primaquine and other 8aminoquinolines particularly effective against the mature gametocytes and the hypnozoites of malaria? Ann Trop Med Parasitol 78:165. 101. Mihaly GW, Ward SA, Edwards G, Orme ML, Breckenridge AM. 1984. Pharmacokinetics of primaquine in man: identification of the carboxylic acid derivative as a major plasma metabolite. Br J Clin Pharmacol 17:441–446. 102. Nasveld PE, Edstein MD, Reid M, Brennan L, Harris IE, Kitchener SJ, Leggat PA, Pickford P, Kerr C, Ohrt C, Prescott W, Tafenoquine Study Team. 2010. Randomized, double-blind study of the safety, tolerability, and efficacy of tafenoquine versus mefloquine for malaria prophylaxis in nonimmune subjects. Antimicrob Agents Chemother 54:792–798. 103. Walsh DS, Wilairatana P, Tang DB, Heppner DG, Jr, Brewer TG, Krudsood S, Silachamroon U, Phumratanaprapin W, Siriyanonda D, Looareesuwan S. 2004. Randomized trial of 3-dose regimens of tafenoquine (WR238605) versus low-dose primaquine for preventing Plasmodium vivax malaria relapse. Clin Infect Dis 39:1095– 1103. 104. Brueckner RP, Coster T, Wesche DL, Shmuklarsky M, Schuster BG. 1998. Prophylaxis of Plasmodium falciparum infection in a human challenge model with WR 238605, a new 8-aminoquinoline antimalarial. Antimicrob Agents Chemother 42:1293–1294. 105. Gu HM, Warhurst DC, Peters W. 1984. Uptake of [3H] dihydroartemisinine by erythrocytes infected with Plasmodium falciparum in vitro. Trans R Soc Trop Med Hyg 78:265–270. 106. Na-Bangchang K, Krudsood S, Silachamroon U, Molunto P, Tasanor O, Chalermrut K, Tangpukdee N, Matangkasombut O, Kano S, Looareesuwan S. 2004. The pharmacokinetics of oral dihydroartemisinin and artesunate in healthy Thai volunteers. Southeast Asian J Trop Med Public Health 35:575–582. 107. Morris CA, Duparc S, Borghini-Fuhrer I, Jung D, Shin CS, Fleckenstein L. 2011. Review of the clinical pharmacokinetics of artesunate and its active metabolite dihydroartemisinin following intravenous, intramuscular, oral or rectal administration. Malar J 10:263. doi:10.1186/ 1475-2875-10-263. 108. Price RN, Nosten F, Luxemburger C, ter Kuile FO, Paiphun L, Chongsuphajaisiddhi T, White NJ. 1996. Effects of artemisinin derivatives on malaria transmissibility. Lancet 347:1654–1658. 109. Chen PQ, Li GQ, Guo XB, He KR, Fu YX, Fu LC, Song YZ. 1994. The infectivity of gametocytes of Plasmodium falciparum from patients treated with artemisinin. Chin Med J (Engl) 107:709–711. 110. Dondorp A, Nosten F, Stepniewska K, Day N, White N. 2005. Artesunate versus quinine for treatment of severe falciparum malaria: a randomised trial. Lancet 366:717– 725. 111. Sinclair D, Donegan S, Isba R, Lalloo DG. 2012. Artesunate versus quinine for treating severe malaria. Cochrane Database Syst Rev (6):CD005967. doi:10.1002/14651858. CD005967.pub4. 112. Dondorp AM, Fanello CI, Hendriksen IC, Gomes E, Seni A, Chhaganlal KD, Bojang K, Olaosebikan R, Anunobi N, Maitland K, Kivaya E, Agbenyega T, Nguah

SB, Evans J, Gesase S, Kahabuka C, Mtove G, Nadjm B, Deen J, Mwanga-Amumpaire J, Nansumba M, Karema C, Umulisa N, Uwimana A, Mokuolu OA, Adedoyin OT, Johnson WB, Tshefu AK, Onyamboko MA, Sakulthaew T, Ngum WP, Silamut K, Stepniewska K, Woodrow CJ, Bethell D, Wills B, Oneko M, Peto TE, von Seidlein L, Day NP, White NJ, AQUAMAT group. 2010. Artesunate versus quinine in the treatment of severe falciparum malaria in African children (AQUAMAT): an open-label, randomised trial. Lancet 376:1647–1657. 113. Talisuna AO, Karema C, Ogutu B, Juma E, Logedi J, Nyandigisi A, Mulenga M, Mbacham WF, Roper C, Guerin PJ, D’Alessandro U, Snow RW. 2012. Mitigating the threat of artemisinin resistance in Africa: improvement of drug-resistance surveillance and response systems. Lancet Infect Dis 12:888–896. 114. Antinori S, Galimberti L, Milazzo L, Corbellino M. 2013. Plasmodium knowlesi: the emerging zoonotic malaria parasite. Acta Trop 125:191–201. 115. Utzinger J, Xiao SH, Tanner M, Keiser J. 2007. Artemisinins for schistosomiasis and beyond. Curr Opin Investig Drugs 8:105–116. 116. Wikman-Jorgensen PE, Henriquez-Camacho CA, Serrano-Villar S, Perez-Molina JA. 2012. The role of artesunate for the treatment of urinary schistosomiasis in schoolchildren: a systematic review and meta-analysis. Pathog Glob Health 106:397–404. 117. Perez del Villar L, Burguillo FJ, Lopez-Aban J, Muro A. 2012. Systematic review and meta-analysis of artemisinin based therapies for the treatment and prevention of schistosomiasis. PLoS One 7:e45867. doi:10.1371/journal.pone.0045867. 118. Nosten F, White NJ. 2007. Artemisinin-based combination treatment of falciparum malaria. Am J Trop Med Hyg 77:181–192. 119. Li Q, Si Y, Xie L, Zhang J, Weina P. 2009. Severe embryolethality of artesunate related to pharmacokinetics following intravenous and intramuscular doses in pregnant rats. Birth Defects Res B Dev Reprod Toxicol 86:385–393. 120. Looareesuwan S, Chulay JD, Canfield CJ, Hutchinson DB, Malarone Clinical Trials Study Group. 1999. Malarone (atovaquone and proguanil hydrochloride): a review of its clinical development for treatment of malaria. Am J Trop Med Hyg 60:533–541. 121. Srivastava IK, Vaidya AB. 1999. A mechanism for the synergistic antimalarial action of atovaquone and proguanil. Antimicrob Agents Chemother 43:1334–1339. 122. Edstein MD, Looareesuwan S, Viravan C, Kyle DE. 1996. Pharmacokinetics of proguanil in malaria patients treated with proguanil plus atovaquone. Southeast Asian J Trop Med Public Health 27:216–220. 123. Radloff PD, Philipps J, Hutchinson D, Kremsner PG. 1996. Atovaquone plus proguanil is an effective treatment for Plasmodium ovale and P. malariae malaria. Trans R Soc Trop Med Hyg 90:682. 124. McAuley JB, Herwaldt BL, Stokes SL, Becher JA, Roberts JM, Michelson MK, Juranek DD. 1992. Diloxanide furoate for treating asymptomatic Entamoeba histolytica cyst passers: 14 years’ experience in the United States. Clin Infect Dis 15:464–468. 125. Markell EK, Udkow MP. 1986. Blastocystis hominis: pathogen or fellow traveler? Am J Trop Med Hyg 35:1023– 1026. 126. Chulay JD, Fleckenstein L, Smith DH. 1988. Pharmacokinetics of antimony during treatment of visceral leishmaniasis with sodium stibogluconate or meglumine antimoniate. Trans R Soc Trop Med Hyg 82:69–72. 127. Oliveira LF, Schubach AO, Martins MM, Passos SL, Oliveira RV, Marzochi MC, Andrade CA. 2011. Systematic review of the adverse effects of cutaneous leishmaniasis treatment in the New World. Acta Trop 118:87–96.

149. Antiparasitic Agents n 128. Murray HW. 2012. Leishmaniasis in the United States: treatment in 2012. Am J Trop Med Hyg 86:434–440. 129. Jha TK, Sundar S, Thakur CP, Bachmann P, Karbwang J, Fischer C, Voss A, Berman J. 1999. Miltefosine, an oral agent, for the treatment of Indian visceral leishmaniasis. N Engl J Med 341:1795–1800. 130. More B, Bhatt H, Kukreja V, Ainapure SS. 2003. Miltefosine: great expectations against visceral leishmaniasis. J Postgrad Med 49:101–103. 131. Prasad R, Kumar R, Jaiswal BP, Singh UK. 2004. Miltefosine: an oral drug for visceral leishmaniasis. Indian J Pediatr 71:143–144. 132. Soto J, Soto P. 2006. Miltefosine: oral treatment of leishmaniasis. Expert Rev Anti Infect Ther 4:177–185. 133. Sindermann H, Croft SL, Engel KR, Bommer W, Eibl HJ, Unger C, Engel J. 2004. Miltefosine (Impavido): the first oral treatment against leishmaniasis. Med Microbiol Immunol 193:173–180. 134. Croft SL, Seifert K, Duchene M. 2003. Antiprotozoal activities of phospholipid analogues. Mol Biochem Parasitol 126:165–172. 135. Bhattacharya SK, Jha TK, Sundar S, Thakur CP, Engel J, Sindermann H, Junge K, Karbwang J, Bryceson AD, Berman JD. 2004. Efficacy and tolerability of miltefosine for childhood visceral leishmaniasis in India. Clin Infect Dis 38:217–221. 136. Sundar S, Jha TK, Thakur CP, Engel J, Sindermann H, Fischer C, Junge K, Bryceson A, Berman J. 2002. Oral miltefosine for Indian visceral leishmaniasis. N Engl J Med 347:1739–1746. 137. Sundar S, Sinha P, Jha TK, Chakravarty J, Rai M, Kumar N, Pandey K, Narain MK, Verma N, Das VN, Das P, Berman J, Arana B. 2013. Oral miltefosine for Indian post-kala-azar dermal leishmaniasis: a randomised trial. Trop Med Int Health 18:96–100. 138. Webster D, Umar I, Kolyvas G, Bilbao J, Guiot MC, Duplisea K, Qvarnstrom Y, Visvesvara GS. 2012. Treatment of granulomatous amoebic encephalitis with voriconazole and miltefosine in an immunocompetent soldier. Am J Trop Med Hyg 87:715–718. 139. Martinez DY, Seas C, Bravo F, Legua P, Ramos C, Cabello AM, Gotuzzo E. 2010. Successful treatment of Balamuthia mandrillaris amoebic infection with extensive neurological and cutaneous involvement. Clin Infect Dis 51:e7–e11. 140. Soto J, Toledo J, Gutierrez P, Nicholls RS, Padilla J, Engel J, Fischer C, Voss A, Berman J. 2001. Treatment of American cutaneous leishmaniasis with miltefosine, an oral agent. Clin Infect Dis 33:E57–E61. 141. Sands M, Kron MA, Brown RB. 1985. Pentamidine: a review. Rev Infect Dis 7:625–634. 142. Waalkes TP, DeVita VT. 1970. The determination of pentamidine (4,4′-diamidinophenoxypentane) in plasma, urine, and tissues. J Lab Clin Med 75:871–878. 143. Davidson RN, den Boer M, Ritmeijer K. 2009. Paromomycin. Trans R Soc Trop Med Hyg 103:653–660. 144. Harris J, Plummer S, Lloyd D. 2001. Antigiardial drugs. Appl Microbiol Biotechnol 57:614–619. 145. Botero D. 1970. Paromomycin as effective treatment of Taenia infections. Am J Trop Med Hyg 19:234–237. 146. Katz M. 1977. Anthelmintics. Drugs 13:124–136. 147. Pepin J, Milord F. 1994. The treatment of human African trypanosomiasis. Adv Parasitol 33:1–47. 148. Dumas M, Bouteille B. 2000. Treatment of human African trypanosomiasis. Bull World Health Organ 78:1474. 149. Taylor HR. 1984. Recent developments in the treatment of onchocerciasis. Bull World Health Organ 62:509–515. 150. Voogd TE, Vansterkenburg EL, Wilting J, Janssen LH. 1993. Recent research on the biological activity of suramin. Pharmacol Rev 45:177–203.

2549

151. Burri C, Baltz T, Giroud C, Doua F, Welker HA, Brun R. 1993. Pharmacokinetic properties of the trypanocidal drug melarsoprol. Chemotherapy 39:225–234. 152. Lutje V, Seixas J, Kennedy A. 2010. Chemotherapy for second-stage human African trypanosomiasis. Cochrane Database Syst Rev (8):CD006201. doi:10.1002/14651858. CD006201.pub2. 153. Kingsnorth AN. 1986. The chemotherapeutic potential of polyamine antimetabolites. Ann R Coll Surg Engl 68:76–81. 154. Huebert ND, Schwartz JJ, Haegele KD. 1997. Analysis of 2-difluoromethyl-DL-ornithine in human plasma, cerebrospinal fluid and urine by cation-exchange high-performance liquid chromatography. J Chromatogr A 762: 293–298. 155. Milord F, Loko L, Ethier L, Mpia B, Pepin J. 1993. Eflornithine concentrations in serum and cerebrospinal fluid of 63 patients treated for Trypanosoma brucei gambiense sleeping sickness. Trans R Soc Trop Med Hyg 87: 473–477. 156. Kennedy PG. 2013. Clinical features, diagnosis, and treatment of human African trypanosomiasis (sleeping sickness). Lancet Neurol 12:186–194. 157. Croft SL. 1997. The current status of antiparasite chemotherapy. Parasitology 114:S3–S15. 158. Fozard JR, Part ML, Prakash NJ, Grove J. 1980. Inhibition of murine embryonic development by alpha-difluoromethylornithine, an irreversible inhibitor of ornithine decarboxylase. Eur J Pharmacol 65:379–391. 159. Polak A, Richle R. 1978. Mode of action of the 2-nitroimidazole derivative benznidazole. Ann Trop Med Parasitol 72:45–54. 160. Murta SM, Ropert C, Alves RO, Gazzinelli RT, Romanha AJ. 1999. In-vivo treatment with benznidazole enhances phagocytosis, parasite destruction and cytokine release by macrophages during infection with a drug-susceptible but not with a derived drug-resistant Trypansoma cruzi population. Parasite Immunol 21:535–544. 161. Docampo R, Moreno SN. 1984. Free radical metabolites in the mode of action of chemotherapeutic agents and phagocytic cells on Trypanosoma cruzi. Rev Infect Dis 6:223–238. 162. Gutteridge WE. 1985. Existing chemotherapy and its limitations. Br Med Bull 41:162–168. 163. Workman P, White RA, Walton MI, Owen LN, Twentyman PR. 1984. Preclinical pharmacokinetics of benznidazole. Br J Cancer 50:291–303. 164. Matta Guedes PM, Gutierrez FR, Nascimento MS, DoValle-Matta MA, Silva JS. 2012. Antiparasitical chemotherapy in Chagas’ disease cardiomyopathy: current evidence. Trop Med Int Health 17:1057–1065. 165. Bern C. 2011. Antitrypanosomal therapy for chronic Chagas’ disease. N Engl J Med 364:2527–2534. 166. Rassi A, Jr, Rassi A, Marin-Neto JA. 2010. Chagas disease. Lancet 375:1388–1402. 167. Jackson Y, Chatelain E, Mauris A, Holst M, Miao Q, Chappuis F, Ndao M. 2013. Serological and parasitological response in chronic Chagas patients 3 years after nifurtimox treatment. BMC Infect Dis 13:85. doi:10.1186/ 1471-2334-13-85. 168. de Toranzo EG, Masana M, Castro JA. 1984. Administration of benznidazole, a chemotherapeutic agent against Chagas disease, to pregnant rats. Covalent binding of reactive metabolites to fetal and maternal proteins. Arch Int Pharmacodyn Ther 272:17–23. 169. Garcia-Bournissen F, Altcheh J, Panchaud A, Ito S. 2010. Is use of nifurtimox for the treatment of Chagas disease compatible with breast feeding? A population pharmacokinetics analysis. Arch Dis Child 95:224–228.

Mechanisms of Resistance to Antiparasitic Agents W. EVAN SECOR, JACQUES LE BRAS, AND JÉRÔME CLAIN

150 Parasitic diseases rank among the most prevalent and severe diseases worldwide and yet their control relies heavily on a single tool: the drugs used for chemotherapy or prophylaxis. This situation exists because no effective antiparasitic vaccines are available and implementation of other control measures often proves to be difficult in countries where parasitic diseases are endemic. This dependence on drugs is compounded by the relative paucity of the current armamentarium of antiparasitic products, a situation attributable largely to a lack of economic incentives for research and development. Furthermore, those drugs that are available are too often used incorrectly in communities and in control programs, a practice that encourages the selection of drugresistant parasites. The complex biologic interactions between parasites and their hosts (and at times vectors) significantly influence the emergence and expression of drug resistance. In many cases the observed resistance is true resistance, attributable to biologic characteristics of the parasites that enable them to survive drug concentrations that are lethal to susceptible members of the species. Mechanisms for such true resistance are varied and include a decrease in drug accumulation within the parasite or modifications in parasite enzyme structure or metabolic pathways. However, various host factors modulate the clinical and parasitologic responses to drug treatment, and the observed responses do not necessarily reflect true parasite resistance or susceptibility. For example, in populations with high rates of exposure to parasitic infections, the resulting high rates of immunity might suffice to eliminate an infection with drug-resistant parasites or treated with an ineffective drug. Conversely, treatment with a drug to which the parasite is biologically susceptible will not necessarily result in therapeutic success if the host takes an inadequate dose of the drug, absorbs it poorly, or lacks the immune response that might be needed for a successful antiparasitic synergism with the drug. Such host factors may be especially important in the areas where most parasitic diseases prevail, where high rates of parasite transmission result in high rates of immunity in most of the population, or where, conversely, malnutrition and human immunodeficiency virus (HIV) infection frequently decrease the patient’s immune status. Greater understanding of the epidemiology and mechanisms of drug resistance can provide valuable guidance for a better use of existing compounds and for the development of novel products. A selective review of drug resistance in five diseases will illustrate the existing problems and their

potential solutions. A summary of the proposed mechanisms of resistance is provided in Table 1.

MALARIA Overview Malaria remains the most visible indicator of the adequacy of health control in all regions of the world with a hot and humid season. By 1955, the World Health Organization (WHO) had established projects for malaria eradication by using indoor residual spraying of insecticides to limit contact between humans and the anophele vector and by mass administration of pyrimethamine and chloroquine to kill erythrocytic-replicating forms of the parasites (1). As these programs faced infrastructure deficiencies and resistance to insecticides, the vector control was often neglected and the bulk of expenditure was devoted to the use of antimalarial drugs against fever, mainly chloroquine, with sulfadoxinepyrimethamine as a secondary drug. Plasmodium falciparum, the most virulent species, has become the dominant species and its resistance to chloroquine and sulfadoxine-pyrimethamine, and further, to all known drugs, has developed to various degrees. Sub-Saharan Africa alone contributes about 80% of the annual 207 million patients worldwide suffering from malaria, leading to an estimated 627,000 deaths (2). Severe malaria affects mainly those without acquired adequate clinical immunity, such as young children or particular groups, such as pregnant women. Since the beginning of the 21st century, a specific effort of integrated control has been made and a reduction in transmission is starting to be observed in parts of Africa (3). The most spectacular action (together with the widespread use of long-lasting insecticidal nets) was the global establishment of malaria treatment with artemisinin-based combination therapies (ACTs), associating a curative dose of a drug with long elimination half-life and a dose of a rapidly active drug able to destroy a large parasite load in few hours. This bitherapy is based on a derivative of artemisinin, a substance extracted from sagebrush grown mainly in China, which generates oxidative stress in the parasite. The combination of multiple drugs enhances clinical efficacy and may delay the acquisition of parasite resistance. We need to consider the history of the emergence and spread of resistance to chloroquine and sulfadoxine-pyrimethamine as premonitory of the risk of losing the effectiveness of ACTs if their use is not controlled.

doi:10.1128/9781555817381.ch150

2550

150. Mechanisms of Resistance to Antiparasitic Agents n TABLE 1

2551

Summary of proposed mechanisms of resistance to selected antiparasitic drugs

Disease Malaria

Drug(s) Chloroquine Pyrimethamine

Sulfadoxine

Atovaquone Artemisinins

Mechanism(s) of resistance Decreased accumulation of the drug by the parasite, resulting from altered transport properties of mutant PfCRT and PfPGH-1 Alteration in binding affinities between the drug and the parasite dihydrofolate reductase-thymidylate synthase, resulting from mutations on the corresponding gene Alteration in binding affinities between the drug and the parasite dihydropteroate synthase, resulting from mutations on the corresponding gene Alteration in binding affinities between the drug and the parasite cytochrome b, resulting from mutations on the corresponding gene Mutated Kelch13 protein; mechanism unknown

Trichomoniasis

Metronidazole, tinidazole

Reduced concentration of enzymes or coenzymes necessary to activate nitro group

Leishmaniasis

Pentavalent antimonials

Decreased active intracellular drug concentration through decreased uptake, increased efflux, or decreased conversion to active trivalent form, glycans that increase host IL-10 production Replaced pentavalent antimonials as primary treatment in some areas, resistance by increased drug efflux or increases in strain infectivity Increased drug efflux, altered thiol metabolism

Miltefosine Amphotericin B African trypanosomiasis

Pentamidine (1st stage) Suramin (1st stage) Melarsoprol (2nd stage) Eflornithine or combination eflornithine/nifurtimox

Schistosomiasis

Praziquantel

Mutation/loss of P2 adenosine and/or aquaglyceroporin 2 transporters that uptake drug Not useful in west Africa where T. b. gambiense is the primary infection because of severe allergic reactions in onchocerciasis patients Mutation/loss of P2 adenosine and/or aquaglyceroporin 2 transporters that uptake drug T. b. rhodesiense naturally tolerant, thus only useful against T. b. gambiense

Widespread clinical resistance not currently recognized as important problem for public health, although genetic bottlenecking observed in some treatment areas and resistance demonstrated in laboratory strains

The 4-amino quinoline drug chloroquine, a cornerstone of antimalarial chemotherapy since the 1940s due to its low cost, its safety, and its rapid action, lost most of its usefulness as the frequency of P. falciparum chloroquine-resistant strains increased and peaked in the 1980s. From limited original foci in Southeast Asia, South America, and Papua New Guinea, resistance has spread inescapably and is now found in most areas of endemicity, including Africa, the continent with the heaviest malaria burden. Chloroquine no longer constitutes an appropriate option for prompt and effective treatment (or prophylaxis) in most countries where P. falciparum malaria is the dominant endemic species. With increased use of sulfadoxine-pyrimethamine, the second affordable, relatively safe, and easily administered drug after chloroquine, parasite resistance to sulfadoxine-pyrimethamine has developed very quickly following the same itineraries as the spread of chloroquine resistance a few years before. Nevertheless, specific groups of people in areas of endemicity still rely on sulfadoxine-pyrimethamine for presumptive treatment of fever or for intermittent preventive treatment as the most effective way to prevent severe consequences of malaria. Losing the two low-cost antimalarials is often seen by experts as a public health disaster. Reducing transmission intensity could slow the spread of resistance but, paradoxically, below a critical level, it may accelerate the selection of multigenic resistance (3). This critical situation has prompted international initiatives to help face the high cost of ACTs, to make affordable rapid-diagnostic

tests available everywhere, and to renew mosquito-control programs by an extensive distribution of insecticide-impregnated nets. Learning from the past that delay in detection and control of resistance to ACTs may ruin all programs without alternatives for decades, forced real-time drug-resistance surveillance to be set up and drug development to be reinforced.

Mechanisms of resistance to selected antimalarials Chloroquine concentrates from nanomolar levels outside the parasite to millimolar levels within the digestive vacuole of the intraerythrocytic trophozoite, where it inhibits hemoglobin degradation (4). Chloroquine forms complexes with hematin, a by-product of host-cell hemoglobin digestion by the parasite, which accumulates in large quantities and eventually kills the parasite. The resistant isolates have in common a defect in chloroquine accumulation in the digestive vacuole (5). Several mechanisms have been proposed to explain the altered chloroquine accumulation, such as changes in the pH gradient or altered membrane permeability leading to a decreased drug uptake or increased drug efflux (6). Chloroquine accumulation has high structural specificity; this suggests the involvement of either a specific transporter/permease or a molecule associated with hematin in the digestive vacuole (7). Following demonstration that chloroquine resistance is reversible by verapamil, earlier studies focused on the orthologue of the mammalian multidrug resistance (mdr) gene, whose products are overex-

2552 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

pressed in cancer cells where they expel cytotoxic drugs (8). This strategy led to the identification of the pfmdr1 gene product (protein PfPGH-1), a transporter located in the membrane of the parasite digestive vacuole (9). Chloroquine susceptibility was found to be altered by PfPGH-1 mutated at specific amino acid residues and in specific parasite genetic backgrounds (10). However, current evidence indicates that pfmdr1 does play a secondary role in chloroquine resistance, except in Madagascar where it plays a major role in chloroquine-treatment failures (11). In following years, using a genetic cross between chloroquinesensitive and chloroquine-resistant parasites, inheritance data led to the key discovery of the P. falciparum chloroquine resistance transporter (pfcrt) gene (12–13). The pfcrt gene encodes a transmembrane protein (PfCRT), as does PfPGH1, located in the membrane of the digestive vacuole (13). A complex set of mutations (or haplotype) of this gene is found in most natural isolates from chloroquine-treatment failures and in isolates with an in vitro chloroquine-resistant phenotype (13–15). Finally, transfection of chloroquinesensitive parasites with the mutant pfcrt genotypes found in resistant isolates suffices to confer chloroquine resistance (16). Mutant PfCRT has acquired the property to expel chloroquine outside of the digestive vacuole (17) which is dependent on a charge loss at codon 76 (usually the K76T mutation) in the first transmembrane helix of the transporter (17–18). The role of the other pfcrt mutations which systematically accompany K76T remains elusive (6). At least four independent mutant pfcrt haplotypes carrying the K76T mutation are seen, varying geographically: Asia-Africa, Papua, and South America 1 and 2 (15). A major event in chloroquine resistance was the emergence on the Thai-Cambodian border in the 1950s of CVIET haplotype in codons 72–76 of the pfcrt gene under drug-pressure selection; this haplotype has now spread to most of the P. falciparum Asian and African territories (15, 19). Nonetheless, the wild pfcrt parasites have not been totally replaced, and their prevalence increases when drug pressure is removed (20). The pfcrt K76T mutation is now a valuable molecular marker used in epidemiological surveys of chloroquine resistance. Its estimated prevalence may offer a useful predictor of the clinical efficacy of chloroquine in a given area, provided that appropriate adjustments are made for host factors, particularly immunity, that may result in parasite clearance in spite of treatment with an ineffective drug (21). Amodiaquine, piperaquine, and pyronaridine, which are in the first line of therapy as partners of artemisinins, share with chloroquine a quinoline scaffold, whereas mefloquine and lumefantrine, the other partners of ACTs, belong to amino-alcohols. Neither the mechanism of action of these drugs nor resistances to them are as clearly understood, as for chloroquine. However, the pfmdr1 gene appears to be involved in parasite response to diverse antimalarials as mutations or amplification of the gene alters parasite susceptibility to desethyl amodiaquine, the active metabolite of amodiaquine (22), quinine, mefloquine, lumefantrine, and artemisinins (10, 23–26). Similarly, wild-type and various mutant pfcrt alleles associate with altered susceptibility to various antimalarials (lumefantrine, amodiaquine, artemisinins) in addition to choloroquine (6, 16, 27). Transporters other than PfCRT and PfPGH-1, such as PfMRP-1 and PfNHE-1, may be involved in resistance to mefloquine, lumefantrine, pyrimethamine (28, 29), and quinine, respectively. Resistance to artemisinins recently emerged in western Cambodia and has now spread across mainland Southeast Asia. It manifests as a delayed parasite-clearance time

following treatment initiation and as higher failure rate (30– 31). Extensive genetic studies have shown that artemisinin resistance is a genetically determined trait (32) and pointed out specific regions of the parasite genome that are associated with the in vivo delayed parasite-clearance time (33– 35). Comparative genomics of laboratory-selected and Cambodian field-derived artemisinin-resistant parasites led to the identification of missense mutations in the kelch13 gene as an in vitro and in vivo marker of artemisinin resistance (36). How this mutant protein contributes to artemisinin resistance remains to be clarified. Regarding drugs that antagonize a single enzyme, such as antifolates or inhibitors of the mitochondrial respiratory chain, a single gene modification is often sufficient to generate high-grade resistance. Malaria parasites mostly rely on de novo synthesis for folate supply. Cycloguanil (produced by the pro-drug proguanil), and its analogue, pyrimethamine, were the first satisfactory synthetic antifolate antimalarials to be on the market in the 1940s. Both inhibit the plasmodial dihydrofolate reductase (PfDHFR), a key enzyme in the folate synthesis pathway of the parasite. Unfortunately, resistance emerged almost instantaneously and independently from several areas where the drugs had been introduced and these antimalarials were soon supplanted by chloroquine (37). A resurgence in the use of PfDHFR inhibitors took place with the demonstration in 1967 that potentiation with other antifolates from the sulfone/sulfonamide group (such as sulfadoxine or dapsone), which inhibit the plasmodial dihydropteroate synthase (PfDHPS), bypassed resistance and sulfadoxine-pyrimethamine became the new spearhead to face chloroquine resistance in Southeast Asia (37). However, resistance to sulfadoxine-pyrimethamine emerged soon after the increased use of sulfadoxine-pyrimethamine in Thailand. Resistance to PfDHFR inhibitors is conferred by mutant PfDHFR enzymes to which antifolate drugs bind less efficiently compared to the wildtype version (38–39). Stepwise acquisitions of pyrimethamine and then sulfadoxine-pyrimethamine resistance in P. falciparum are mirrored by the stepwise acquisitions of PfDHFR mutations: first the key mutation S108N, then the additive mutations N51I and C59R, and finally the additive I164L mutation (39, 40). Other additive mutations can be found in specific areas such as in South America (41). The quadruple-mutant N51I-C59R-S108N-I164L which associates with the highest level of sulfadoxine-pyrimethamine resistance as well as to decreased sensitivity to chloproguanil-dapsone (an attempt to develop a new antifolate combination similar to sulfadoxine-pyrimethamine) has been reported mostly in Southeast Asia. However, the triple mutant N51I-C59R-S108N, which is widespread in Africa, also associates with treatment failure to sulfadoxine-pyrimethamine (42). This triple mutant N51I-C59R-S108N pfdhfr gene emerged in Southeast Asia, and spread all over Asia and Africa in the following years (43, 44). Resistance to sulfone/sulfonamide group (sulfadoxine being the major antimalarial compound) is conferred by mutant PfDHPS enzymes to which antifolate drugs bind less efficiently compared to the wild-type version (45, 46). Resistance to sulfadoxine has been traced to a set of sequential mutations in the pfdhps gene. The likely initial event consisted of an A437G mutation, with subsequent additional mutations conferring increasing degrees of resistance (37). These resistance mutations have appeared independently multiple times and in multiple endemic sites (47). Altogether, resistance to the sulfadoxine-pyrimethamine combination appears to require three mutations in the pfdhfr

150. Mechanisms of Resistance to Antiparasitic Agents n

gene, and the probability of sulfadoxine-pyrimethamine treatment failure increases with additional mutations in pfdhps. The clinical outcome of a sulfadoxine-pyrimethamine treatment is subjected to additional host factors such as the level of folates and of acquired immunity and drug absorption and metabolism (42, 48). Cross-resistance has been demonstrated between cycloguanil and pyrimethamine (38, 39, 49). Consequently, interest in other antifolates, such as chlorproguanil plus dapsone (LapDap), to treat parasites resistant to sulfadoxine-pyrimethamine has been limited. The combination of atovaquone and proguanil was registered in 1996 in North America and Europe where, within 10 years, it became the most used antimalarial for prophylaxis and first-line treatment of non-severe P. falciparum malaria. Atovaquone is a ubiquinone analogue and binds to cytochrome b (PfCytB), a component of the complex III in the mitochondrial-respiratory chain (50). In association with proguanil, the effective concentration at which it collapses the mitochondrial membrane potential is diminished (51). In addition, proguanil is partially metabolized to the antifolate cycloguanil by human P450 cytochromes. The contribution of the resulting low cycloguanil blood concentrations to the therapeutic efficacy of atovaquone-proguanil remains yet to be substantiated. As with the antifolates, atovaquone resistance emerged almost instantaneously (52). Resistance to atovaquone is conferred by mutant PfCytB to which atovaquone binds less efficiently compared to the wild-type version (53). The substitution of Y for S or N or C in codon 268 of PfCytB is found associated with treatment failures and confers a high level of atovaquone resistance (54) that proguanil could not thwart (55). Remarkably, the resistance mutation is not detected in areas of endemicity (56), and it seems to evolve repeatedly during primary infections (57). Due to the risk of rapid extension of resistance and high cost of the drug, the deployment of atovaquone in regions where malaria is endemic has not been considered as a priority. Chloroquine resistance in Plasmodium vivax, the second most common malaria parasite, has been very limited despite widespread chloroquine use. P. vivax, which relapses from dormant parasites in the liver, developed partial resistance to primaquine, the only drug active against liver forms (58). Primaquine’s diminished efficacy is also associated with polymorphisms in the host cytochrome P-450 2D6, resulting in altered concentrations of the active metabolites (59). While drug resistance in P. vivax remains at low magnitude, increasing attention has recently focused to tackle this threat. Investigations on mechanisms of resistance in this species currently examine the potential role of P. vivax homologues of pfcrt (60), pfmdr1 (61), pfdhfr (62), and pfdhps (63).

TRICHOMONIASIS Infection with Trichomonas vaginalis is one of the most common causes of human vaginitis as well as the most prevalent nonviral sexually transmitted disease (64). T. vaginalis infections are associated with preterm delivery, low birth weight, and greater susceptibility to infection with HIV as well as increased shedding of virus in HIV-infected individuals (65– 67). As a result, expedient treatment of this infection has become an important public health concern (68, 69). T. vaginalis is a facultative anaerobe and trichomoniasis is most commonly treated with the 5-nitroimidazole class of drugs. Two members of this group, metronidazole and tinidazole, are the only drugs licensed for treatment of

2553

trichomoniasis in the United States. Tinidazole is more active at equimolar concentrations than metronidazole and is recommended if treatment with metronidazole fails (70– 72). However, strains clinically resistant to metronidazole can have cross-resistance to tinidazole. The molecular epidemiology of T. vaginalis suggests that clinically resistant isolates are genetically related and are concentrated within one of two major subpopulations (73–75). In a survey of women attending sexually transmitted disease clinics in six U.S. cities, 4.3% of isolates exhibited drug resistance (76), suggesting that almost 160,000 residents in the U.S. alone may be affected (171). The 5-nitroimidazoles enter parasites in an inactive form by passive diffusion. Once inside, the drug is reduced to the active nitro-radical anion that is thought to cause parasite death by breaking or disrupting DNA. A number of electron donors have been proposed for drug activation, including ferredoxin, pyruvate-ferredoxin oxidoreductase (PFOR), and malic enzyme in the hydrogenosome, which is the source of ATP generation in these amitochondriate parasites (77– 79). Drug resistance occurs when transcription of one or more of these enzymes is decreased, and laboratory-generated resistant isolates have smaller hydrogenosomes (80). However, clinical resistance does not correlate with lower transcript levels of these enzymes or smaller hydrogenesome size (80, 81). The nitroimidazole drugs are also reduced by the flavin enzyme thioredoxin reductase and covalently bind and inhibit proteins associated with thioredoxin-mediated redox regulation (82). An in vitro-induced nitroimidazole-resistant strain demonstrated reduced thioredoxin reductase activity, not as a result of decreased enzyme concentration but because of a deficiency in the necessary FAD cofactor. Trichomonads with minimal PFOR and malate-dehydrogenase activity remained susceptible to metronidazole (82). Furthermore, use of a flavin inhibitor rendered a normally susceptible isolate resistant to high concentrations of metronidazole. Flavin inhibition also reduced levels of PFOR and malate dehydrogenase, suggesting that the decreased hydrogenosomal redox enzymes observed in resistant isolates may be a consequence rather than a cause of metronidazole resistance (83). A third possible mechanism for nitroimidazole resistance was suggested by the observation that trichomonas isolates that harbored Mycoplasma hominis symbionts had a mean in vitro resistance level 10-fold higher than noninfected trichomonads (84). However, the increased mean resistance of the M. hominis-infected isolates was still lower than typically observed for resistant isolates. In a separate study, there was no association of mycoplasma infection with clinical resistance (85). It remains unclear which, if any, of these mechanisms is responsible for the clinical nitroimidazole resistance observed in some T. vaginalis infections. Treatment of patients who have metronidazole-resistant trichomoniasis often results in an immediate resolution of symptoms and a negative wet mount. However, within 3 to 4 weeks, in the absence of further exposure, symptoms may recur as the number of organisms rises. Thus, it is important to monitor efficacy of treatment for up to a month and to encourage patients to avoid unprotected intercourse during this time. When nitroimidazole resistance is encountered, patients are often successfully treated with increased doses of drug for a longer time (71, 72); however, many patients cannot tolerate high doses of metronidazole and such practices may only exacerbate the development of drug resistance. In addition, some patients experience hypersensitivity reactions in response to metronidazole and tinidazole.

2554 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

Clearly, alternatives to the nitroimidazoles for treatment of T. vaginalis infections are needed (86). A library of 1,040 FDA-approved drugs was recently tested against metronidazole-sensitive and -resistant isolates in vitro in an attempt to find alternate oral therapies for trichomoniasis. While no drugs were as effective as the 5-nitroimidazoles, some enhanced the activity of metronidazole when used in combination (87). Intravaginal treatment with drugs such as furazolidone and paromomycin sulfate that are not absorbed well from the intestine or cannot be ingested has been successful to cure some patients but, in general, has limited efficacy (86, 88). Povidone-iodine and boric acid have also shown efficacy for some patients but additional clinical testing is needed (88, 89).

LEISHMANIASIS Leishmaniasis is transmitted to humans by phlebotomine sandfly vectors and manifests itself in a variety of syndromes. Depending in part on which of the possible 20 species of Leishmania that infects humans is present, pathology can range from a cutaneous lesion that is self-limiting to the more severe mucosal or visceral forms. The identification of leishmaniasis as an important opportunistic infection in patients with AIDS has presented new challenges for treatment of this disease, with increased treatment failures and drug toxicity in HIV-1 positive individuals (90, 91). Use of antileishmanial drugs is limited by their high cost, the difficulty of their administration (injections for several weeks), and/or their associated toxicity. These factors are even more consequential in the developing countries where leishmaniasis is endemic and can lead to premature selftermination of therapy, which in turn may promote increased levels of resistance (92). While true drug resistance has been described for isolates of some Leishmania spp., other species or isolates may just differ in their intrinsic sensitivity to certain compounds (93–96). True resistance is more likely in anthroponotic forms of leishmaniaisis, such as Leishmania donovani and Leishmania tropica, because the zoonotic species that primarily infect animals, with humans as an occasional host, rarely encounter drugs and serve as a reservoir for drug-sensitive parasites (94, 97). The frontline drugs for treating Leishmania infections caused by all species and all clinical forms have long been the pentavalent-antimonial compounds such as sodium stibogluconate and meglumine antimoniate. These compounds are inexpensive compared to other anti-leishmanial drugs but their extensive use has led to widespread treatment failure. For example, in some areas of endemicity in India, treatment failure of visceral leishmaniasis caused by L. donovani is as high as 65% (98). Evidence for true drug resistance in this setting comes from observations that isolates from clinically resistant patients require higher in vitro concentrations of drug to kill the parasites than do isolates from patients who respond to treatment (99). However, the correlation between clinical outcome of treatment for leishmaniasis and the in vitro susceptibility of the causative isolate is not always clear (93–95, 100). Pentavalent antimonials are prodrugs that become reduced within the mammalian host cell or parasite to an active trivalent form that boosts the intracellular cytotoxic potential of macrophages, as well as disrupts the parasite’s redox metabolism; many of the proposed mechanisms of resistance across the various Leishmania spp. involve a lower intracellular concentration of active drug (101, 102). One mechanism by which this occurs is a decrease in aquaglyceroporins on the parasite’s surface, resulting in less uptake of

drug (103, 104). Resistance has also been associated with increased production of trypanothione or glutathione that binds with the trivalent antimonials and sequestration of the resulting thiol-drug conjugates into intracellular organelles by ATP-binding cassette (ABC) transporters (101, 105, 106). Increased expression of the enzymes involved in thiol synthesis or of the ABC transporter promotes resistance to pentavalent antimonials; inhibition of these pathways in resistant strains increases susceptibility to the drug (102, 103, 105–109). Adding to the difficulty of defining resistance mechanisms, more than one of these mechanisms may arise, even within closely related parasite strains (101). Genomic, proteomic, and metabolomic approaches have further revealed the complex, multifactorial nature of antimonial resistance in Leishmania spp. as well as indicate other mechanisms that could be in play, such as protection from reactive-oxygen species and stress response (95, 96, 100, 101, 104, 110, 111). Because of the high level of pentavalent-antimonial resistance, this treatment is no longer considered useful in parts of India and, in these places, miltefosine is now the primary treatment for visceral leishmaniasis (96, 112). Miltefosine was originally developed as an anti-cancer drug and is particularly promising because it can be taken orally and has fewer side effects than most of the parenteral treatments for leishmaniasis (113). As a result, although the drug is more expensive than other therapies, it can be administered on an outpatient basis, reducing the overall cost for treatment in comparison to less costly drugs that require hospitalization. Enthusiasm is tempered, however, both because it is teratogenic, thereby limiting its unregulated use in women of child-bearing age, as well as the observation that leishmanial resistance to miltefosine develops easily in vitro (113). Relapse rates of 20% within 1 year after miltefosine treatment have already been reported (114). As with the antimonials, resistance to miltefosine has been associated with mechanisms that decrease intracellular concentrations, and inhibitors of ABC transporters can restore drug susceptibility in vitro (115–117). However, recent studies suggest that these mechanisms do not explain all of the clinical resistance to miltefosine and other mechanisms, such as infectivity of the parasite strain, may be involved (112, 114, 118). Another drug now being used for primary treatment of visceral leishmaniasis is amphotericin B, which interacts with parasite-specific 24-alkyl sterols and induces pore formation in the parasite plasma membranes (92). The use of amphotericin B has been limited in the past due to its high cost and toxicity; however, new lipid-associated formulations of amphotericin B have greatly reduced toxicities and retain good efficacy even when administered in lower doses (119). Like the pentavalent antimonials, optimal activity of amphotericin B may require competent host immune responses (120, 121). Nevertheless, amphotericin B seems to be superior to antimonials for treatment of leishmaniasis in HIV-infected individuals (90, 91). Lipid-associated formulations of amphotericin B are phagocytized by host monocytes and accumulate in the phagocytic lysosomes where Leishmania amastigotes reside. Although parasite isolates from HIV-positive patients who relapsed with L. infantum demonstrated no decrease in their in vitro susceptibility to amphotericin B (121), clinically resistant isolates have been obtained from patients infected with L. donovani. Interestingly, although the mode of action of amphotericin B is thought to differ greatly from that of the antimonials, field isolates indicated that those with greater sodium-antimonygluconate resistance had greater in vitro resistance to ampho-

150. Mechanisms of Resistance to Antiparasitic Agents n

tericin B (115). As with the other anti-lesihmania drugs, amphotericin B resistance is associated with greater drug efflux and altered thiol metabolism (122). Part of the challenge for understanding drug activity and treatment failure during leishmaniasis relates to the intracellular location of the amastigotes within host macrophages. As a result, treatment efficacy involves host factors in addition to the parasites and therapeutic compounds (123). For example, intracellular killing of amastigotes is dependent on macrophage production of nitric oxide, which is inhibited by the cytokine interleukin-10 (IL-10). Interestingly, some antimony-resistant isolates of L. donovani express a unique glycan that upregulates the ABC transporter in the parasite as well as IL-10 production by the host (108, 124). As a result, neutralization of IL-10 has been suggested as an adjunct therapy for visceral leishmaniasis (125).

2555

discovery of a high-affinity pentamidine transporter (HAPT) that also concentrates pentamidine within the parasites (132). Recently, HAPT has been identified as aquaglyceroporin 2, and loss of this receptor is shown to confer melarsoprol/pentamidine cross-resistance (133, 134). T. brucei parasites also express an ABC transporter that, similar to Leishmania, functions as an efflux pump and may contribute to the melarsoprol resistance of certain isolates (127, 128, 135). While melarsoprol treatment failure may be as high as 20% in some settings (136), field resistance to eflornithine and nifurtimox has not been documented. Nevertheless, resistance in laboratory strains has been readily induced for both drugs (137, 138). Eflornithine resistance is associated with the loss of a nonessential amino acid transporter that is responsible for drug uptake, raising the concern that field resistance could easily develop (137).

AFRICAN TRYPANOSOMIASIS SCHISTOSOMIASIS Trypanosoma brucei rhodesiense and Trypanosoma brucei gambiense are the etiologic agents of human African trypanosomiasis (HAT). The two subspecies are endemic in eastand west-central Africa, respectively, with T. b. gambiense causing the vast majority of infections (126, 127). Because these parasites possess antigenic-switching mechanisms, host immune responses are ineffective and the prospects for the development of vaccines against these organisms are meager. Control efforts have largely been focused on prevention through reduction of the tsetse fly vector, usually through traps or insecticides (126). However, recent political unrest with subsequent loss of an effective public health infrastructure in parts of Africa has caused a resurgence of this disease. As a result, drug treatment is the only medical intervention available to combat sleeping sickness for the foreseeable future. HAT has two stages; an initial bloodstream stage followed by invasion of the central nervous system in the second stage, which causes the meningoencephalitic systems associated with “sleeping sickness,” the more familiar name for HAT (126). Pentamidine and suramin are drugs used for treatment of the first stage but not advanced disease because they are highly ionic and do not cross the bloodbrain barrier. Furthermore, use of suramin is avoided for treatment of T. b. gambiense because of the risk of severe allergic reactions that it can cause in onchocerciasis patients (126, 127). While clinical resistance to these drugs does not seem to be a problem, failures can occur when infections are diagnosed and treated after disease has progressed past the hemolymphatic stage. Late-stage, central nervous system disease is treated with melarsoprol, eflornithine, or nifurtimox-eflornitihine combination therapy (126, 128). Eflornithine is effective against late-stage T. b. gambiense infections that are resistant to melarsoprol, but T. b. rhodesiense parasites are naturally tolerant to this drug (129). The drugs for HAT are difficult to administer or are relatively toxic, which can contribute to premature cessation of treatment that can in turn contribute to the development of drug resistance. Pentamidine and melarsoprol share an amidinium-like moiety with amino purines that is recognized and actively taken up by nucleoside transporters in the trypanosome membrane, resulting in concentration of these compounds within the parasite (127). One of these receptors, the T. brucei P2 adenosine transporter, or TbAT1, has been extensively studied and resistance to pentamidine and melarsoprol is associated with loss of functional tbat1 expression, including in field isolates (127, 130, 131). However, tbat1 knockout parasites were only partially resistant, leading to the

Drug resistance in human helminths is rare, a fact attributed to their long reproduction cycles and to their lack of multiplication inside the human host (the exception being Strongyloides stercoralis). Thus, when considering treatment failures in schistosomiasis, it is important to distinguish characteristics leading to reduced drug efficacy from true drug resistance. For example, persons with very high levels of infection are less likely to be cured with single-dose therapy than individuals with lower worm burdens (139, 140). This is in part related to the fact that the standard drug used to treat schistosomiasis, praziquantel, is only effective against the adult stage of the parasite and, even under the best conditions, a single dose of praziquantel does not demonstrate complete efficacy (141, 142). Immature worms that may be present at the time of drug treatment, especially in areas of high transmission, are not susceptible to the initial treatment and subsequently develop into patent infections that give the impression of drug resistance. As a result, two treatments spaced 4 to 6 weeks apart are more effective than a single treatment and should be attempted when drug resistance is suspected (139, 143, 144). Rapid reinfection in areas of high transmission should also be considered as a possible explanation for apparent praziquantel-treatment failure (145, 146). Suspected true praziquantel resistance has been described for Schistosoma mansoni infections in Egypt and Kenya (147, 148). Eggs obtained from the feces of individuals who could not be successfully treated have been used to establish infections in mice, confirming the drug-resistance phenotype (147–149). However, widespread clinical resistance has not developed, even in areas with high-intensity treatment pressure for a prolonged time (150–152). Laboratory and field studies have shown decreased diversity of schistosomes following praziquantel treatment suggestive of a genetic bottleneck, a warning sign for development of drug resistance (153, 154). However, this observation has not been consistent in all field studies (155, 156). The exact mechanism of praziquantel, and as a result the mechanism of drug resistance, is not definitively understood. In drug-susceptible parasites, praziquantel-induced damage to the tegument of adult schistosomes renders the worms susceptible to attack and killing by the host’s immune response; the effect of drug on resistant parasites is reduced (157). The unique beta subunit of the schistosome calcium-ion channel is a molecular target for praziquantel with treatment rapidly inducing a calcium-dependent, sustained muscle contraction in the worm’s tegument (158, 159). However, no differences in this gene’s sequence or expression were observed among a limited number of praziquantel-resistant and -susceptible

2556 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

parasite strains (160). Furthermore, cytochalasin D reverses the effects of praziquantel without altering the calcium influx, thus raising the possibility that this is not the effector of parasite death (161). Other proposed mechanisms of praziquantel action include inhibition of the worm’s nucleoside uptake and binding to, and altering the function of, schistosome myosin-light chain (162, 163). One potential mechanism of drug resistance includes increased expression of a P-glycoprotein ATP-dependent efflux-pump homologue in parasite strains that have reduced susceptibility to praziquantel (164, 165). Cross-breeding of adult worms with different levels of sensitivity to drug results in offspring with an intermediate phenotype, suggesting that at least artificially induced resistance displays partial dominance (166). Differential gene transcription between susceptible and resisitant parasites and between adult and juvenile worms in response to praziquantel may also play a role in drug susceptibility (167, 168). Although widespread praziquantel resistance has not been observed, as mass drug-administration programs are increasingly employed, there is a fear that resistance may emerge (169). This risk reiterates the need for ongoing monitoring of praziquantel resistance and efforts for the discovery of new drugs or repositioning of existing compounds to treat for schistosomiasis (170).

FUTURE PERSPECTIVES Several factors contribute to the emergence of drug-resistant parasites. Those parasite species with short life cycles and high multiplication rates that occur in areas of intense transmission are most likely to develop resistant subpopulations. The selection of such populations is encouraged when the parasites are repeatedly exposed to suboptimal drug concentrations. This pattern can result from the use of drugs with long half-lives, or more typically, from the frequent, often unjustified use, of inadequate doses of drugs, a common occurrence in countries where parasite infections are endemic. Public health interventions to correct these factors have not always been successful and would benefit from a better understanding of the drug-resistance mechanisms used by parasites. These mechanisms are very diverse and have been difficult to study, but recent technological advances now provide long-awaited tools that will facilitate the task. The genome sequences for several Plasmodium species, T. vaginalis, Leishmania major, T. brucei, S. mansoni, and Schistosoma haematobium, have been compiled. When these data are used, for example, in combination with microarray technology or whole-genome sequencing, where the DNA of drug-resistant parasite strains is compared to drug-susceptible strains, identification of the genes that confer resistance should proceed even more rapidly than in the last few years. In addition, genomic, transcriptomic, and proteomic data may also be useful in the design of new chemotherapeutic agents as they help researchers identify metabolic processes of parasites that are sufficiently different from those of their human hosts to allow specific attack.

REFERENCES 1. World Health Organization. 1993. A global strategy for malaria control. World Health Organization, Geneva, Switzerland. 2. World Health Organization. 2014. World malaria report 2013. World Health Organization, Geneva, Switzerland. 3. Talisuna AO, Bloland P, D’Alessandro U. 2004. History, dynamics and public health importance of malaria parasite resistance. Clin Microbiol Rev 17:235–254.

4. Fitch CD. 1970. Plasmodium falciparum in owl monkeys: drug resistance and chloroquine binding capacity. Science 169:289–290. 5. Verdier F, Le Bras J, Clavier F, Hatin I, Blayo MC. 1985. Chloroquine uptake by Plasmodium falciparum-infected human erythrocytes during in vitro culture and its relationship to chloroquine resistance. Antimicrob Agents Chemother 27:561–564. 6. Ecker A, Lehane AM, Clain J, Fidock DA. 2012. PfCRT and its role in antimalarial drug resistance. Trends Parasitol 28:504–514. 7. Bray PG, Mungthin M, Ridley RG, Ward SA. 1998. Access to hematin: the basis of chloroquine resistance. Mol Pharmacol 54:170–179. 8. Martin SK, Oduola AM, Milhous WK. 1987. Reversal of chloroquine resistance in Plasmodium falciparum by verapamil. Science 235:899–901. 9. Foote SJ, Thompson JK, Cowman AF, Kemp DJ. 1989. Amplification of the multidrug resistance gene in some chloroquine-resistant isolates of P. falciparum. Cell 57:921– 930. 10. Reed MB, Saliba KJ, Caruana SR, Kirk K, Cowman AF. 2000. Pgh1 modulates sensitivity and resistance to multiple antimalarials in Plasmodium falciparum. Nature 403:906– 909. 11. Andriantsoanirina V, Ratsimbasoa A, Bouchier C, Tichit M, Jahevitra M, Rabearimanana S, Raherinjafy R, Mercereau-Puijalon O, Durand R, Ménard D. 2010. Chloroquine clinical failures in P. falciparum malaria are associated with mutant Pfmdr-1, not Pfcrt in Madagascar. PLoS One 5:e13281. doi:10.1371/journal.pone.0013281 12. Su X, Kirkman LA, Fujioka H, Wellems TE. 1997. Complex polymorphisms in an approximately 330 kDa protein are linked to chloroquine-resistant P. falciparum in Southeast Asia and Africa. Cell 91:593–603. 13. Fidock DA, Nomura T, Talley AK, Cooper RA, Dzekunov SM, Ferdig MT, Ursos LM, Sidhu AB, Naudé B, Deitsch KW, Su XZ, Wootton JC, Roepe PD, Wellems TE. 2000. Mutations in the P. falciparum digestive vacuole transmembrane protein PfCRT and evidence for their role in chloroquine resistance. Mol Cell 6:861–871. 14. Djimdé A, Doumbo OK, Cortese JF, Kayentao K, Doumbo S, Diourté Y, Dicko A, Su XZ, Nomura T, Fidock DA, Wellems TE, Plowe CV. 2001. A molecular marker for chloroquine-resistant falciparum malaria. N Eng J Med 344:257–263. 15. Wootton JC, Feng X, Ferdig MT, Cooper RA, Mu J, Baruch DI, Magill AJ, Su XZ. 2002. Genetic diversity and chloroquine selective sweeps in Plasmodium falciparum. Nature 418:320–323. 16. Sidhu AB, Verdier-Pinard D, Fidock DA. 2002. Chloroquine resistance in Plasmodium falciparum malaria parasites conferred by pfcrt mutations. Science 298:210–213. 17. Martin RE, Marchetti RV, Cowan AI, Howitt SM, Bröer S, Kirk K. 2009. Chloroquine transport via the malaria parasite’s chloroquine resistance transporter. Science 325: 1680–1682. 18. Johnson DJ, Fidock DA, Mungthin M, Lakshmanan V, Sidhu AB, Bray PG, Ward SA. 2004. Evidence for a central role for PfCRT in conferring Plasmodium falciparum resistance to diverse antimalarial agents. Mol Cell 15:867– 877. 19. Ariey F, Fandeur T, Durand R, Randrianarivelojosia M, Jambou R, Legrand E, Ekala MT, Bouchier C, Cojean S, Duchemin JB, Robert V, Le Bras J, Mercereau-Puijalon O. 2006. Invasion of Africa by a single pfcrt allele of South East Asian type. Malar J 5:34. 20. Kublin JG, Cortese JF, Njunju EM, Mukadam RA, Wirima JJ, Kazembe PN, Djimdé AA, Kouriba B, Taylor TE, Plowe CV. 2003. Reemergence of chloroquine-sensitive

150. Mechanisms of Resistance to Antiparasitic Agents n

21.

22.

23.

24.

25.

26.

27.

28.

29.

30.

31.

32.

33.

Plasmodium falciparum malaria after cessation of chloroquine use in Malawi. J Infect Dis 187:1870–1875. Djimdé A, Doumbo OK, Steketee RW, Plowe CV. 2001. Application of a molecular marker for surveillance of chloroquine-resistant falciparum malaria. Lancet 358:890–891. Holmgren G, Gil JP, Ferreira PM, Veiga MI, Obonyo CO, Björkman A. 2006. Amodiaquine resistant Plasmodium falciparum malaria in vivo is associated with selection of pfcrt 76T and pfmdr1 86Y. Infect Genet Evol 6:309–314. Price RN, Uhlemann AC, Brockman A, McGready R, Ashley E, Phaipun L, Patel R, Laing K, Looareesuwan S, White NJ, Nosten F, Krishna S. 2004. Mefloquine resistance in Plasmodium falciparum and increased pfmdr1 gene copy number. Lancet 364:438–447. Sidhu AB, Valderramos SG, Fidock DA. 2005. pfmdr1 mutations contribute to quinine resistance and enhance mefloquine and artemisinin sensitivity in Plasmodium falciparum. Mol Microbiol 57:913–926. Sisowath C, Strömberg J, Mårtensson A, Msellem M, Obondo C, Björkman A, Gil JP. 2005. In vivo selection of Plasmodium falciparum pfmdr1 86N coding alleles by artemether-lumefantrine (Coartem). J Infect Dis 191:1014– 1017. Sidhu AB, Uhlemann AC, Valderramos SG, Valderramos JC, Krishna S, Fidock DA. 2006. Decreasing pfmdr1 copy number in Plasmodium falciparum malaria heightens susceptibility to mefloquine, lumefantrine, halofantrine, quinine, and artemisinin. J Infect Dis 194:528–535. Sisowath C, Petersen I, Veiga MI, Mårtensson A, Premji Z, Björkman A, Fidock DA, Gil JP. 2009. In vivo selection of Plasmodium falciparum parasites carrying the chloroquine-susceptible pfcrt K76 allele after treatment with artemether-lumefantrine in Africa. J Infect Dis 199:750– 757. Dahlström S, Ferreira PE, Veiga MI, Sedighi N, Wiklund L, Mårtensson A, Färnert A, Sisowath C, Osório L, Darban H, Andersson B, Kaneko A, Conseil G, Björkman A, Gil JP. 2009. Plasmodium falciparum multidrug resistance protein 1 and artemisinin-based combination therapy in Africa. J Infect Dis 200:1456–1464. Dahlström S, Veiga MI, Mårtensson A, Björkman A, Gil JP. 2009. Polymorphism in PfMRP1 (Plasmodium falciparum multidrug resistance protein 1) amino acid 1466 associated with resistance to sulfadoxine-pyrimethamine treatment. Antimicrob Agents Chemother 53:2553–2556. Dondorp AM, Nosten F, Yi P, Das D, Phyo AP, Tarning J, Lwin KM, Ariey F, Hanpithakpong W, Lee SJ, Ringwald P, Silamut K, Imwong M, Chotivanich K, Lim P, Herdman T, An SS, Yeung S, Singhasivanon P, Day NP, Lindegardh N, Socheat D, White NJ. 2009. Artemisinin resistance in Plasmodium falciparum malaria. N Engl J Med 361:455–467. Carrara VI, Lwin KM, Phyo AP, Ashley E, Wiladphaingern J, Sriprawat K, Rijken M, Boel M, McGready R, Proux S, Chu C, Singhasivanon P, White N, Nosten F. 2013. Malaria burden and artemisinin resistance in the mobile and migrant population on the Thai-Myanmar border, 1999–2011: an observational study. PLoS Med 10:e1001398. doi:10.1371/journal.pmed.1001398 Phyo AP, Nkhoma S, Stepniewska K, Ashley EA, Nair S, McGready R, ler Moo C, Al-Saai S, Dondorp AM, Lwin KM, Singhasivanon P, Day NP, White NJ, Anderson TJ, Nosten F. 2012. Emergence of artemisinin-resistant malaria on the western border of Thailand: a longitudinal study. Lancet 379:1960–1966. KCheeseman IH, Miller BA, Nair S, Nkhoma S, Tan A, T a n J C, Al Sa ai S, Ph yo AP , M oo CL , L win KM, McGready R, Ashley E, Imwong M, Stepniewska K, Yi P, Dondorp AM, Mayxay M, Newton PN, White NJ, Nosten F, Ferdig MT, Anderson TJ. 2012. A major genome region

34.

35.

36.

37.

38.

39.

40.

41.

42.

43.

44.

2557

underlying artemisinin resistance in malaria. Science 336: 79–82. Takala-Harrison S, Clark TG, Jacob CG, Cummings MP, Miotto O, Dondorp AM, Fukuda MM, Nosten F, Noedl H, Imwong M, Bethell D, Se Y, Lon C, Tyner SD, Saunders DL, Socheat D, Ariey F, Phyo AP, Starzengruber P, Fuehrer HP, Swoboda P, Stepniewska K, Flegg J, Arze C, Cerqueira GC, Silva JC, Ricklefs SM, Porcella SF, Stephens RM, Adams M, Kenefic LJ, Campino S, Auburn S, MacInnis B, Kwiatkowski DP, Su XZ, White NJ, Ringwald P, Plowe CV. 2013. Genetic loci associated with delayed clearance of Plasmodium falciparum following artemisinin treatment in Southeast Asia. Proc Natl Acad Sci USA 110:240–245. Miotto O, Almagro-Garcia J, Manske M, Macinnis B, Campino S, Rockett KA, Amaratunga C, Lim P, Suon S, Sreng S, Anderson JM, Duong S, Nguon C, Chuor CM, Saunders D, Se Y, Lon C, Fukuda MM, AmengaEtego L, Hodgson AV, Asoala V, Imwong M, TakalaHarrison S, Nosten F, Su XZ, Ringwald P, Ariey F, Dolecek C, Hien TT, Boni MF, Thai CQ, AmambuaNgwa A, Conway DJ, Djimdé AA, Doumbo OK, Zongo I, Ouedraogo JB, Alcock D, Drury E, Auburn S, Koch O, Sanders M, Hubbart C, Maslen G, Ruano-Rubio V, Jyothi D, Miles A, O’Brien J, Gamble C, Oyola SO, Rayner JC, Newbold CI, Berriman M, Spencer CC, McVean G, Day NP, White NJ, Bethell D, Dondorp AM, Plowe CV, Fairhurst RM, Kwiatkowski DP. 2013. Multiple populations of artemisinin-resistant Plasmodium falciparum in Cambodia. Nat Genet 45:648–655. Ariey F, Witkowski B, Amaratunga C, Beghain J, Langlois AC, Khim N, Kim S, Duru V, Bouchier C, Ma L, Lim P, Leang R, Duong S, Sreng S, Suon S, Chuor CM, Bout DM, Ménard S, Rogers WO, Genton B, Fandeur T, Miotto O, Ringwald P, Le Bras J, Berry A, Barale JC, Fairhurst RM, Benoit-Vical F, Mercereau-Puijalon O, Ménard D. 2014. A molecular marker of artemisininresistant Plasmodium falciparum malaria. Nature 505:50– 55. Gregson A, Plowe CV. 2005. Mechanisms of resistance of malaria parasites to antifolates. Pharmacol Rev 57:117– 145. Peterson DS, Walliker D, Wellems TE. 1988. Evidence that a point mutation in dihydrofolate reductase-thymidylate synthase confers resistance to pyrimethamine in falciparum malaria. Proc Natl Acad Sci USA 85:9114–9118. Sirawaraporn W, Sathitkul T, Sirawaraporn R, Yuthavong Y, Santi DV. 1997. Antifolate-resistant mutants of Plasmodium falciparum dihydrofolate reductase. Proc Natl Acad Sci USA 94:1124–1129. Lozovsky ER, Chookajorn T, Brown KM, Imwong M, Shaw PJ, Kamchonwongpaisan S, Neafsey DE, Weinreich DM, Hartl DL. 2009. Stepwise acquisition of pyrimethamine resistance in the malaria parasite. Proc Natl Acad Sci USA 106:12025–12030. Cortese JF, Caraballo A, Contreras CE, Plowe CV. 2002. Origin and dissemination of Plasmodium falciparum drugresistance mutations in South America. J Infect Dis 186: 999–1006. Plowe CV, Cortese JF, Djimde A, Nwanyanwu OC, Watkins WM, Winstanley PA, Estrada-Franco JG, Mollinedo RE, Avila JC, Cespedes JL, Carter D, Doumbo OK. 1997. Mutations in Plasmodium falciparum dihydrofolate reductase and dihydropteroate synthase and epidemiologic patterns of pyrimethamine-sulfadoxine use and resistance. J Infect Dis 176:1590–1596. Roper C, Pearce R, Nair S, Sharp B, Nosten F, Anderson T. 2004. Intercontinental spread of pyrimethamine-resistant malaria. Science 305:1124. Maïga O, Djimdé AA, Hubert V, Renard E, Aubouy A, Kironde F, Nsimba B, Koram K, Doumbo OK, Le Bras

2558 n

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

J, Clain J. 2007. A shared Asian origin of the triple mutant dhfr allele in P. falciparum from sites across Africa. J Infect Dis. 196:165–172. Wang P, Read M, Sims PF, Hyde JE. 1997. Sulfadoxine resistance in the human malaria parasite Plasmodium falciparum is determined by mutations in dihydropteroate synthetase and an additional factor associated with folate utilization. Mol Microbiol 23:979–986. Triglia T, Wang P, Sims PF, Hyde JE, Cowman AF. 1998. Allellic exchange at the endogenous genomic locus in Plasmodium falciparum proves the role of dihydropteroate synthase in sulfadoxine-resistant malaria. EMBO J 17: 3807–3815. Pearce RJ, Pota H, Evehe MS, Bâ el-H, Mombo-Ngoma G, Malisa AL, Ord R, Inojosa W, Matondo A, Diallo DA, Mbacham W, van den Broek IV, Swarthout TD, Getachew A, Dejene S, Grobusch MP, Njie F, Dunyo S, Kweku M, Owusu-Agyei S, Chandramohan D, Bonnet M, Guthmann JP, Clarke S, Barnes KI, Streat E, Katokele ST, Uusiku P, Agboghoroma CO, Elegba OY, Cissé B, A-Elbasit IE, Giha HA, Kachur SP, Lynch C, Rwakimari JB, Chanda P, Hawela M, Sharp B, Naidoo I, Roper C. 2009. Multiple origins and regional dispersal of resistant dhps in African Plasmodium falciparum malaria. PLoS Med 6:e1000055. doi:10.1371/journal.pmed.1000055 Kublin JG, Dzinjalamala FK, Kamwendo DD, Malkin EM, Cortese JF, Martino LM, Mukadam RA, Rogerson SJ, Lescano AG, Molyneux ME, Winstanley PA, Chimpeni P, Taylor TE, Plowe CV. 2002. Molecular markers for failure of sulfadoxine-pyrimethamine and chlorproguanil-dapsone treatment of Plasmodium falciparum malaria. J Infect Dis 185:380–388. Basco LK, Ringwald P. 2000. Molecular epidemiology of malaria in Yaounde, Cameroon. VI. Sequence variations in the Plasmodium falciparum dihydrofolate reductase-thymidylate synthase gene and in vitro resistance to pyrimethamine and cycloguanil. Am J Trop Med Hyg 62:271–276. Srivastava IK, Morrisey JM, Darrouzet E, Daldal F, Vaidya AB. 1999. Resistance mutations reveal the atovaquone-binding domain of cytochrome b in malaria parasites. Mol Microbiol 33:704–711. Srivastava IK, Vaidya AB. 1999. A mechanism for the synergistic antimalarial action of atovaquone and proguanil. Antimicrob Agents Chemother 43:1334–1339. Looareesuwan SC, Viravan C, Webster HK, Kyle DE, Hutchinson DB, Canfield CJ. 1996. Clinical studies on atovaquone, alone or in association with other antimalarial drugs, for treatment of acute uncomplicated malaria in Thailand. Am J Trop Med Hyg 54:62–66. Kessl JJ, Ha KH, Merritt AK, Lange BB, Hill P, Meunier B, Meshnick SR, Trumpower BL. 2005. Cytochrome b mutations that modify the ubiquinol-binding pocket of the cytochrome bc1 complex and confer anti-malarial drug resistance in Saccharomyces cerevisiae. J Biol Chem 280:17142–17148. Musset L, Bouchaud O, Matheron S, Massias L, Le Bras J. 2006. Clinical atovaquone-proguanil resistance of Plasmodium falciparum associated with cytochrome b codon 268 mutations. Microbes Infect 8:2599–2604. Fivelman QL, Adagu IS, Warhurst DC. 2004. Modified fixed-ratio isobologram method for studying in vitro interactions between atovaquone and proguanil or dihydroartemisinin against drug-resistant strains of Plasmodium falciparum. Antimicrob Agents Chemother 48:4097–4102. Musset L, Pradines B, Parzy D, Durand R, Bigot P, Le Bras J. 2006. Apparent absence of atovaquone/proguanil resistance in 477 Plasmodium falciparum isolates from untreated French travellers. J Antimicrob Chemother 57:110– 115. Musset L, Le Bras J, Clain J. 2007. Parallel evolution of adaptive mutations in Plasmodium falciparum mitochondrial

58.

59.

60.

61.

62.

63.

64.

65.

66.

67.

68. 69.

70.

71.

72.

73.

DNA during atovaquone-proguanil treatment. Mol Biol Evol 24:1582–1585. Smoak BL, DeFraites RF, Magill AJ, Kain KC, Wellde BT. 1997. Plasmodium vivax infections in U.S. Army troops: failure of primaquine to prevent relapse in studies from Somalia. Am J Trop Med Hyg 56:231–234. Bennett JW, Pybus BS, Yadava A, Tosh D, Sousa JC, McCarthy WF, Deye G, Melendez V, Ockenhouse CF. 2013. Primaquine failure and cytochrome P-450 2D6 in Plasmodium vivax malaria. N Engl J Med 369:1381–1382. Nomura T, Carlton JM, Baird JK, del Portillo HA, Fryauff DJ, Rathore D, Fidock DA, Su X, Collins WE, McCutchan TF, Wootton JC, Wellems TE. 2001. Evidence for different mechanisms of chloroquine resistance in 2 Plasmodium species that cause human malaria. J Infect Dis 183:1653–1661. Brega S, Meslin B, de Montbrison F, Severini C, Gradoni L, Udomsangpetch R, Sutanto I, Peyron F, Picot S. 2005. Identification of the Plasmodium vivax mdr-like gene (pvmdr1) and analysis of single-nucleotide polymorphisms among isolates from different areas of endemicity. J Infect Dis 191:272–277. Hastings MD, Porter KM, Maguire JD, Susanti I, Kania W, Bangs MJ, Sibley CH, Baird JK. 2004. Dihydrofolate reductase mutations in Plasmodium vivax from Indonesia and therapeutic response to sulfadoxine plus pyrimethamine. J Infect Dis 189:744–750. Korsinczky M, Fischer K, Chen N, Baker J, Rieckmann K, Cheng Q. 2004. Sulfadoxine resistance in Plasmodium vivax is associated with a specific amino acid in dihydropteroate synthase at the putative sulfadoxine-binding site. Antimicrob Agents Chemother 48:2214–2222. World Health Organization. 2011. Prevalence and incidence of selected sexually transmitted infections, Chlamydia trachomatis, Neisseria gonorrhoeae, syphilis and Trichomonas vaginalis: methods and results used by WHO to generate 2005 estimates. http://whqlibdoc.who.int/publi cations/2011/9789241502450_eng.pdf. Mann JR, McDermott S, Gill T. 2010. Sexually transmitted infection is associated with increased risk of preterm birth in South Carolina women insured by Medicaid. J Matern Fetal Neonatal Med 23:563–568. Van Der Pol B, Kwok C, Pierre-Louis B, Rinaldi A, Salata RA, Chen PL, van de Wijgert J, Mmiro F, Mugerwa R, Chipato T, Morrison CS. 2008. Trichomonas vaginalis infection and human immunodeficiency virus acquisition in African women. J Infect Dis 197:548–554. Kissinger P, Amedee A, Clark RA, Dumestre J, Theall KP, Myers L, Hagensee ME, Farley TA, Martin DH. 2009. Trichomonas vaginalis treatment reduces vaginal HIV1 shedding. Sex Transm Dis 36:11–16. McClelland RS. 2008. Trichomonas vaginalis infection: can we afford to do nothing? J Infect Dis 197:487–489. van der Pol B. 2007. Trichomonas vaginalis infection: the most prevalent nonviral sexually transmitted infection receives the least public health attention. Clin Infect Dis 44:23–25. Crowell AL, Sanders-Lewis KA, Secor WE. 2003. In vitro metronidazole and tinidazole activities against metronidazole-resistant strains of Trichomonas vaginalis. Antimicrob Agents Chemother 47:1407–1409. Workowski KA, Berman SM. 2011. Centers for Disease Control and Prevention Sexually Transmitted Diseases Treatment Guidelines. Clin Infect Dis 53(Suppl 3):S59– S63. Bosserman EA, Helms DJ, Mosure DJ, Secor WE, Workowski KA. 2011. Utility of antimicrobial susceptibility testing in Trichomonas vaginalis-infected women with clinical treatment failure. Sex Transm Dis 38:983–987. Snipes LJ, Gamard PM, Narcisi EM, Beard CB, Lehmann T, Secor WE. 2000. Molecular epidemiology of metronida-

150. Mechanisms of Resistance to Antiparasitic Agents n

74.

75.

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

zole resistance in a population of Trichomonas vaginalis clinical isolates. J Clin Microbiol 38:3004–3009. Conrad MD, Gorman AW, Schillinger JA, Fiori PL, Arroyo R, Malla N, Dubey ML, Gonzalez J, Blank S, Secor WE, Carlton JM. 2012. Extensive genetic diversity, unique population structure and evidence of genetic exchange in the sexually transmitted parasite Trichomonas vaginalis. PLoS Negl Trop Dis 6:e1573. doi:10.1371/journal.pntd. 0001573 Paulish-Miller TE, Augostini P, Schuyler JA, Smith WL, Mordechai E, Adelson ME, Gygax SE, Secor WE, Hilbert DW. 2014. Trichomonas vaginalis metronidazole resistance is associated with single nucleotide polymorphisms in the nitroreductase genes ntr4Tv and ntr6Tv. Antimicrob Agents Chemother 58:2938–2943. Kirkcaldy RD, Augostini P, Asbel LE, Bernstein KT, Kerani RP, Mettenbrink CJ, Pathela P, Schwebke JR, Secor WE, Workowski KA, Davis D, Braxton J, Weinstock HS. 2012. Trichomonas vaginalis antimicrobial drug resistance in 6 US cities, STD Surveillance Network, 2009–2010. Emerg Infect Dis 18:939–943. Land KM, Delgadillo MG, Johnson PJ. 2002. In vivo expression of ferredoxin in a drug resistant trichomonad increases metronidazole susceptibility. Mol Biochem Parasitol 121:153–157. Rasoloson D, Vanácová S, Tomková E, Rázga J, Hrdy I, Tachezý J, Kulda J. 2002. Mechanisms of in vitro development of resistance to metronidazole in Trichomonas vaginalis. Microbiology 148:2467–2477. Hrdý I, Cammack R, Stopka P, Kulda J, Tachezy J. 2005. Alternative pathway of metronidazole activation in Trichomonas vaginalis hydrogenosomes. Antimicrob Agents Chemother 49:5033–5036. Wright JM, Webb RI, O’Donoghue P, Upcroft P, Upcroft JA. 2010. Hydrogenosomes of laboratory-induced metronidazole-resistant Trichomonas vaginalis lines are downsized while those from clinically metronidazole-resistant isolates are not. J Eukaryot Microbiol 57:171–176. Mead JR, Fernadez M, Romagnoli PA, Secor WE. 2006. Use of Trichomonas vaginalis clinical isolates to evaluate correlation of gene expression and metronidazole resistance. J Parasitol 92:196–199. Leitsch D, Kolarich D, Binder M, Stadlmann J, Altmann F, Duchêne M. 2009. Trichomonas vaginalis: metronidazole and other nitroimidazole drugs are reduced by the flavin enzyme thioredoxin reductase and disrupt the cellular redox system. Implications for nitroimidazole toxicity and resistance. Mol Microbiol 72:518–536. Leitsch D, Kolarich D, Duchêne M. 2010. The flavin inhibitor diphenyleneiodonium renders Trichomonas vaginalis resistant to metronidazole, inhibits thioredoxin reductase and flavin reductase, and shuts off hydrogenosomal enzymatic pathways. Mol Biochem Parasitol 171:17–24. Xiao JC, Xie LF, Fang SL, Gao MY, Zhu Y, Song LY, Zhong HM, Lun ZR. 2006. Symbiosis of Mycoplasma hominis in Trichomonas vaginalis may link metronidazole resistance in vitro. Parasitol Res 100:123–130. Butler SE, Augostini P, Secor WE. 2010. Mycoplasma hominis infection of Trichomonas vaginalis is not associated with metronidazole-resistant trichomoniasis in clinical isolates from the United States. Parasitol Res 107:1023–1027. Helms DJ, Mosure DJ, Secor WE, Workowski KA. 2008. Management of Trichomonas vaginalis in women with suspected metronidazole hypersensitivity. Am J Obstet Gynecol 198:370.e1–7. Goodhew EB, Secor WE. 2013. Drug library screening against metronidazole-sensitive and metronidazole-resistant Trichomonas vaginalis isolates. Sex Transm Infect 89:479–484.

2559

88. Waters LJ, Dave SS, Deayton JR, French PD. 2005. Recalcitrant Trichomonas vaginalis infection—a case series. Int J STD AIDS 16:505–509. 89. Muzny C, Barnes A, Mena L. 2012. Symptomatic Trichomonas vaginalis infection in the setting of severe nitroimidazoleallergy:successfultreatmentwithboricacid.SexHealth 9:389–391. 90. Jarvis JN, Lockwood DN. 2013. Clinical aspects of visceral leishmaniasis in HIV infection. Curr Opin Infect Dis 26:1–9. 91. Cota GF, de Sousa MR, Fereguetti TO, Rabello A. 2013. Efficacy of anti-leishmania therapy in visceral leishmaniasis among HIV infected patients: a systematic review with indirect comparison. PLoS Negl Trop Dis 7:e2195. doi:10 .1371/journal.pntd.0002195 92. Mishra J, Saxena A, Singh S. 2007. Chemotherapy of leishmaniasis: past, present and future. Curr Med Chem 14:1153–1169. 93. Rijal S, Yardley V, Chappuis F, Decuypere S, Khanal B, Singh R, Boelaert M, De Doncker S, Croft S, Dujardin JC. 2007. Antimonial treatment of visceral leishmaniasis: are current in vitro susceptibility assays adequate for prognosis of in vivo therapy outcome? Microbes Infect 9: 529–535. 94. Aït-Oudhia K, Gazanion E, Vergnes B, Oury B, Sereno D. 2011. Leishmania antimony resistance: what we know what we can learn from the field. Parasitol Res 109:1225– 1232. 95. Torres DC, Ribeiro-Alves M, Romero GA, Dávila AM, Cupolillo E. 2013. Assessment of drug resistance related genes as candidate markers for treatment outcome prediction of cutaneous leishmaniasis in Brazil. Acta Trop 126: 132–141. 96. Berg M, Mannaert A, Vanaerschot M, Van Der Auwera G, Dujardin JC. 2013. (Post-) Genomic approaches to tackle drug resistance in Leishmania. Parasitology 140: 1492–1505. 97. Natera S, Machuca C, Padrón-Nieves M, Romero A, Díaz E, Ponte-Sucre A. 2007. Leishmania spp.: proficiency of drug-resistant parasites. Int J Antimicrob Agents 29:637– 642. 98. Croft SL, Sundar S, Fairlamb AH. 2006. Drug resistance in leishmaniasis. Clin Microbiol Rev 19:111–126. 99. Sundar S. 2001. Drug resistance in Indian visceral leishmaniasis. Trop Med Int Health 6:849–854. 100. Adaui V, Maes I, Huyse T, Van den Broeck F, Talledo M, Kuhls K, De Doncker S, Maes L, Llanos-Cuentas A, Schönian G, Arevalo J, Dujardin JC. 2011. Multilocus genotyping reveals a polyphyletic pattern among naturally antimony-resistant Leishmania braziliensis isolates from Peru. Infect Genet Evol 11:1873–1880. 101. Decuypere S, Vanaerschot M, Brunker K, Imamura H, Müller S, Khanal B, Rijal S, Dujardin JC, Coombs GH. 2012. Molecular mechanisms of drug resistance in natural Leishmania populations vary with genetic background. PLoS Negl Trop Dis 6:e1514. doi:10.1371/journal.pntd .0001514 102. do Monte-Neto RL, Coelho AC, Raymond F, Légaré D, Corbeil J, Melo MN, Frézard F, Ouellette M. 2011. Gene expression profiling and molecular characterization of antimony resistance in Leishmania amazonensis. PLoS Negl Trop Dis 5:e1167. doi:10.1371/journal.pntd.0001167 103. Rai S, Bhaskar, Goel SK, Nath Dwivedi U, Sundar S, Goyal N. 2013. Role of efflux pumps and intracellular thiols in natural antimony resistant isolates of Leishmania donovani. PLoS One 8:e74862. doi:10.1371/journal.pone. 0074862 104. Mukherjee A, Boisvert S, do Monte-Neto RL, Coelho AC, Raymond F, Mukhopadhyay R, Corbeil J, Ouellette M. 2013. Telomeric gene deletion and intrachromosomal

2560 n

105.

106.

107.

108.

109.

110.

111.

112.

113.

114.

115.

116.

117.

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

amplification in antimony-resistant Leishmania. Mol Microbiol 88:189–202. Mittal MK, Rai S, Ashutosh, Ravinder, Gupta S, Sundar S, Goyal N. 2007. Characterization of natural antimony resistance in Leishmania donovani isolates. Am J Trop Med Hyg 76:681–688. Mukherjee A, Padmanabhan PK, Singh S, Roy G, Girard I, Chatterjee M, Ouellette M, Madhubala R. 2007. Role of ABC transporter MRPA, γ-glutamylcysteine synthetase and ornithine decarboxylase in natural antimonyresistant isolates of Leishmania donovani. J Antimicrob Chemother 59:204–211. Walker J, Gongora R, Vasquez JJ, Drummelsmith J, Burchmore R, Roy G, Ouellette M, Gomez MA, Saravia NG. 2012. Discovery of factors linked to antimony resistance in Leishmania panamensis through differential proteome analysis. Mol Biochem Parasitol 183:166–176. Mukhopadhyay R, Mukherjee S, Mukherjee B, Naskar K, Mondal D, Decuypere S, Ostyn B, Prajapati VK, Sundar S, Dujardin JC, Roy S. 2011. Characterisation of antimony-resistant Leishmania donovani isolates: biochemical and biophysical studies and interaction with host cells. Int J Parasitol 41:1311–1321. Manzano JI, García-Hernández R, Castanys S, Gamarro F. 2013. A new ABC half-transporter in Leishmania major is involved in resistance to antimony. Antimicrob Agents Chemother 57:3719–3730. Matrangolo FS, Liarte DB, Andrade LC, de Melo MF, Andrade JM, Ferreira RF, Santiago AS, Pirovani CP, Silva-Pereira RA, Murta SM. 2013. Comparative proteomic analysis of antimony-resistant and -susceptible Leishmania braziliensis and Leishmania infantum chagasi lines. Mol Biochem Parasitol 190:63–75. Berg M, Vanaerschot M, Jankevics A, Cuypers B, Maes I, Mukherjee S, Khanal B, Rijal S, Roy S, Opperdoes F, Breitling R, Dujardin JC. 2013. Metabolic adaptations of Leishmania donovani in relation to differentiation, drug resistance, and drug pressure. Mol Microbiol 90:428–442. Bhandari V, Kulshrestha A, Deep DK, Stark O, Prajapati VK, Ramesh V, Sundar S, Schonian G, Dujardin JC, Salotra P. 2012. Drug susceptibility in Leishmania isolates following miltefosine treatment in cases of visceral leishmaniasis and post kala-azar dermal leishmaniasis. PLoS Negl Trop Dis 6:e1657. doi:10.1371/journal.pntd. 0001657 Sundar S, Olliaro PL. 2007. Miltefosine in the treatment of leishmaniasis: clinical evidence for informed clinical risk management. Ther Clin Risk Manag 3:733–740. Rijal S, Ostyn B, Uranw S, Rai K, Bhattarai NR, Dorlo TP, Beijnen JH, Vanaerschot M, Decuypere S, Dhakal SS, Das ML, Karki P, Singh R, Boelaert M, Dujardin JC. 2013. Increasing failure of miltefosine in the treatment of Kala-azar in Nepal and the potential role of parasite drug resistance, reinfection, or noncompliance. Clin Infect Dis 56:1530–1538. Kumar D, Kulshrestha A, Singh R, Salotra P. 2009. In vitro susceptibility of field isolates of Leishmania donovani to miltefosine and amphotericin B: correlation with sodium antimony gluconate susceptibility and implications for treatment in areas of endemicity. Antimicrob Agents Chemother 53:835–838. Sánchez-Cañete MP, Carvalho L, Pérez-Victoria FJ, Gamarro F, Castanys S. 2009. Low plasma membrane expression of the miltefosine transport complex renders Leishmania braziliensis refractory to the drug. Antimicrob Agents Chemother 53:1305–1313. Pérez-Victoria JM, Bavchvarov BI, Torrecillas IR, Martínez-García M, López-Martín C, Campillo M, Castanys S, Gamarro F. 2011. Sitamaquine overcomes ABC-mediated resistance to miltefosine and antimony in Leishmania. Antimicrob Agents Chemother 55:3838–3854.

118. Rai K, Cuypers B, Bhattarai NR, Uranw S, Berg M, Ostyn B, Dujardin JC, Rijal S, Vanaerschot M. 2013. Relapse after treatment with miltefosine for visceral leishmaniasis is associated with increased infectivity of the infecting Leishmania donovani strain. MBio 4:e00611–13. 119. Bern C, Adler-Moore J, Berenguer J, Boelaert M, den Boer M, Davidson RN, Figueras C, Gradoni L, Kafetzis DA, Ritmeijer K, Rosenthal E, Royce C, Russo R, Sundar S, Alvar J. 2006. Liposomal amphotericin B for the treatment of visceral leishmaniasis. Clin Infect Dis 43:917–924. 120. Mueller M, Ritmeijer K, Balasegaram M, Koummuki Y, Santana MR, Davidson R. 2007. Unresponsiveness to AmBisome in some Sudanese patients with kala-azar. Trans R Soc Trop Med Hyg 101:19–24. 121. Lachaud L, Bourgeois N, Plourde M, Leprohon P, Bastien P, Ouellette M. 2009. Parasite susceptibility to amphotericin B in failures of treatment for visceral leishmaniasis in patients coinfected with HIV type 1 and Leishmania infantum. Clin Infect Dis 48:e16–22. 122. Purkait B, Kumar A, Nandi N, Sardar AH, Das S, Kumar S, Pandey K, Ravidas V, Kumar M, De T, Singh D, Das P. 2012. Mechanism of amphotericin B resistance in clinical isolates of Leishmania donovani. Antimicrob Agents Chemother 56:1031–1041. 123. Rijal S, Bhandari S, Koirala S, Singh R, Khanal B, Loutan L, Dujardin JC, Boelaert M, Chappuis F. 2010. Clinical risk factors for therapeutic failure in kala-azar patients treated with pentavalent antimonials in Nepal. Trans R. Soc Trop Med Hyg 104:225–229. 124. Mukherjee B, Mukhopadhyay R, Bannerjee B, Chowdhury S, Mukherjee S, Naskar K, Allam US, Chakravortty D, Sundar S, Dujardin JC, Roy S. 2013. Antimonyresistant but not antimony-sensitive Leishmania donovani up-regulates host IL-10 to overexpress multidrug-resistant protein 1. Proc Natl Acad Sci USA 110:E575–582. 125. Gautam S, Kumar R, Maurya R, Nylén S, Ansari N, Rai M, Sundar S, Sacks D. 2011. IL-10 neutralization promotes parasite clearance in splenic aspirate cells from patients with visceral leishmaniasis. J Infect Dis 204: 1134–1137. 126. Brun R, Blum J, Chappuis F, Burri C. 2010. Human African trypanosomiasis. Lancet 375:148–159. 127. Baker N, de Koning HP, Mäser P, Horn D. 2013. Drug resistance in African trypanosomiasis: the melarsoprol and pentamidine story. Trends Parasitol 29:110–118. 128. Barrett MP, Vincent IM, Burchmore RJ, Kazibwe AJ, Matovu E. 2011. Drug resistance in human African trypanosomiasis. Future Microbiol 6:1037–1047. 129. Delespaux V, de Koning HP. 2007. Drugs and drug resistance in African trypanosomiasis. Drug Resist Updat 10:30–50. 130. Stewart ML, Burchmore RJ, Clucas C, Hertz-Fowler C, Brooks K, Tait A, Macleod A, Turner CM, De Koning HP, Wong PE, Barrett MP. 2010. Multiple genetic mechanisms lead to loss of functional TbAT1 expression in drug-resistant trypanosomes. Eukaryot Cell 9:336–343. 131. Nerima B, Matovu E, Lubega GW, Enyaru JC. 2007. Detection of mutant P2 adenosine transporter (TbAT1) gene in Trypanosoma brucei gambiense isolates from northwest Uganda using allele-specific polymerase chain reaction. Trop Med Int Health 12:1361–1368. 132. de Koning HP. 2008. Ever-increasing complexities of diamidine and arsenical crossresistance in African trypanosomes. Trends Parasitol 24:345–349. 133. Baker N, Glover L, Munday JC, Aguinaga Andrés D, Barrett MP, de Koning HP, Horn D. 2012. Aquaglyceroporin 2 controls susceptibility to melarsoprol and pentamidine in African trypanosomes. Proc Natl Acad Sci USA 109:10996–11001.

150. Mechanisms of Resistance to Antiparasitic Agents n 134. Munday JC, Eze AA, Baker N, Glover L, Clucas C, Aguinaga Andrés D, Natto MJ, Teka IA, McDonald J, Lee RS, Graf FE, Ludin P, Burchmore RJ, Turner CM, Tait A, Macleod A, Mäser P, Barrett MP, Horn D, De Koning HP. 2013. Trypanosoma brucei aquaglyceroporin 2 is a high-affinity transporter for pentamidine and melaminophenyl arsenic drugs and the main genetic determinant of resistance to these drugs. J Antimicrob Chemother 69:651–663. 135. Lüscher A, Nerima B, Mäser P. 2006. Combined contributions of TbAT1 and TbMRPA to drug resistance in Trypanosoma brucei. Mol Biochem Parasitol 150:364–366. 136. Robays J, Nyamowala G, Sese C, Betu Ku Mesu Kande V, Lutumba P, Van der Veken W, Boelaert M. 2008. High failure rates of melarsoprol for sleeping sickness, Democratic Republic of Congo. Emerg Infect Dis 14:966– 967. 137. Vincent IM, Creek D, Watson DG, Kamleh MA, Woods DJ, Wong PE, Burchmore RJ, Barrett MP. 2010. A molecular mechanism for eflornithine resistance in African trypanosomes. PLoS Pathog 6:e1001204. doi:10.1371/ journal.ppat.1001204 138. Sokolova AY, Wyllie S, Patterson S, Oza SL, Read KD, Fairlamb AH. 2010. Cross-resistance to nitro drugs and implications for treatment of human African trypanosomiasis. Antimicrob Agents Chemother 54:2893–2900. 139. Utzinger J, N’Goran EK, N’Dri A, Lengeler C, Tanner M. 2000. Efficacy of praziquantel against Schistosoma mansoni with particular consideration for intensity of infection. Trop Med Intl Health 5:771–778. 140. Montresor A. 2011. Cure rate is not a valid indicator for assessing drug efficacy and impact of preventive chemotherapy interventions against schistosomiasis and soiltransmitted helminthiasis. Trans R Soc Trop Med Hyg 105:361–363. 141. Tchuem Tchuenté LA, Momo SC, Stothard JR, Rollinson D. 2013. Efficacy of praziquantel and reinfection patterns in single and mixed infection foci for intestinal and urogenital schistosomiasis in Cameroon. Acta Trop 128:275–283. 142. Reta B, Erko B. 2013. Efficacy and side effects of praziquantel in the treatment for Schistosoma mansoni infection in school children in Senbete Town, northeastern Ethiopia. Trop Med Int Health 18:1338–1343. 143. Tukahebwa EM, Vennervald BJ, Nuwaha F, Kabatereine NB, Magnussen P. 2013. Comparative efficacy of one versus two doses of praziquantel on cure rate of Schistosoma mansoni infection and re-infection in Mayuge District, Uganda. Trans R Soc Trop Med Hyg 107:397–404. 144. King CH, Olbrych SK, Soon M, Singer ME, Carter J, Colley DG. 2011. Utility of repeated praziquantel dosing in the treatment of schistosomiasis in high-risk communities in Africa: a systematic review. PLoS Negl Trop Dis 5:e1321. doi:10.1371/journal.pntd.0001321 145. Garba A, Lamine MS, Barkiré N, Djibo A, Sofo B, Gouvras AN, Labbo R, Sebangou H, Webster JP, Fenwick A, Utzinger J. 2013. Efficacy and safety of two closely spaced doses of praziquantel against Schistosoma haematobium and S. mansoni and re-infection patterns in school-aged children in Niger. Acta Trop 128:334–344. 146. Webster BL, Diaw OT, Seye MM, Faye DS, Stothard JR, Sousa-Figueiredo JC, Rollinson D. 2013. Praziquantel treatment of school children from single and mixed infection foci of intestinal and urogenital schistosomiasis along the Senegal River Basin: monitoring treatment success and re-infection patterns. Acta Trop 128:292–302. 147. Melman SD, Steinauer ML, Cunningham C, Kubatko LS, Mwangi IN, Wynn NB, Mutuku MW, Karanja DM, Colley DG, Black CL, Secor WE, Mkoji GM, Loker ES. 2009. Reduced susceptibility to praziquantel among

148.

149.

150.

151.

152.

153.

154.

155.

156.

157.

158.

159.

160.

161.

162.

2561

naturally occurring Kenyan isolates of Schistosoma mansoni. PLoS Negl Trop Dis 3:e504. doi:10.1371/journal.pntd .0000504 Ismail M, Metwally A, Farghaly A, Bruce J, Tao LF, Bennett JL. 1996. Characterization of isolates of Schistosoma mansoni from Egyptian villagers that tolerate high doses of praziquantel. Am J Trop Med Hyg 55:214–218. Lamberton PH, Hogan SC, Kabatereine NB, Fenwick A, Webster JP. 2010. In vitro praziquantel test capable of detecting reduced in vivo efficacy in Schistosoma mansoni human infections. Am J Trop Med Hyg 83:1340–1347. Black CL, Steinauer ML, Mwinzi PN, Secor W, Karanja DM, Colley DG. 2009. Impact of intense, longitudinal retreatment with praziquantel on cure rates of Schistosomiasis mansoni in a cohort of occupationally exposed adults in western Kenya. Trop Med Int Health 14:450–457. Guidi A, Andolina C, Makame Ame S, Albonico M, Cioli D, Juma Haji H. 2010. Praziquantel efficacy and long-term appraisal of schistosomiasis control in Pemba Island. Trop Med Int Health 15:614–618. Seto EY, Wong BK, Lu D, Zhong B. 2011. Human schistosomiasis resistance to praziquantel in China: should we be worried? Am J Trop Med Hyg 85:74–82. Coeli R, Baba EH, Araujo N, Coelho PM, Oliveira G. 2013. Praziquantel treatment decreases Schistosoma mansoni genetic diversity in experimental infections. PLoS Negl Trop Dis 7:e2596. doi:10.1371/journalpntd.0002596 Norton AJ, Gower CM, Lamberton PH, Webster BL, Lwambo NJ, Blair L, Fenwick A, Webster JP. 2010. Genetic consequences of mass human chemotherapy for Schistosoma mansoni: population structure pre- and postpraziquantel treatment in Tanzania. Am J Trop Med Hyg 83:951–957. Blanton RE, Blank WA, Costa JM, Carmo TM, Reis EA, Silva LK, Barbosa LM, Test MR, Reis MG. 2011. Schistosoma mansoni population structure and persistence after praziquantel treatment in two villages of Bahia, Brazil. Int J Parasitol 41:1093–1099. Huyse T, Van den Broeck F, Jombart T, Webster BL, Diaw O, Volckaert FA, Balloux F, Rollinson D, Polman K. 2013. Regular treatments of praziquantel do not impact on the genetic make-up of Schistosoma mansoni in Northern Senegal. Infect Genet Evol 18:100–105. Doenhoff MJ, Cioli D, Utzinger J. 2008. Praziquantel: mechanisms of action, resistance and new derivatives for schistosomiasis. Curr Opin Infect Dis 21:659–667. Nogi T, Zhang D, Chan JD, Marchant JS. 2009. A novel biological activity of praziquantel requiring voltageoperated Ca2+ channel β subunits: subversion of flatworm regenerative polarity. PLoS Negl Trop Dis 3:e464. doi:10. 1371/journal.pntd.0000464 Greenberg RM. 2013. New approaches for understanding mechanisms of drug resistance in schistosomes. Parasitology 140:1534–1546. Valle C, Troiani AR, Festucci A, Pica-Mattoccia L, Liberti P, Wolstenholme A, Francklow K, Doenhoff MJ, Cioli D. 2003. Sequence and level of endogenous expression of calcium channel β subunits in Schistosoma mansoni displaying different susceptibilities to praziquantel. Mol Biochem Parasitol 130:111–115. Pica-Mattoccia L, Orsini T, Basso A, Festucci A, Liberti P, Guidi A, Marcatto-Maggi AL, Nobre-Santana S, Troiani AR, Cioli D, Valle C. 2008. Schistosoma mansoni: lack of correlation between praziquantel-induced intraworm calcium influx and parasite death. Exp Parasitol 119:332–335. Angelucci F, Basso A, Bellelli A, Brunori M, Pica Mattoccia L, Valle C. 2007. The anti-schistosomal drug praziquantel is an adenosine antagonist. Parasitology 134:1215–1221.

2562 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

163. Gnanasekar M, Salunkhe AM, Mallia AK, He YX, Kalyanasundaram R. 2009. Praziquantel affects the regulatory myosin light chain of Schistosoma mansoni. Antimicrob Agents Chemother 53:1054–1060. 164. Messerli SM, Kasinathan RS, Morgan W, Spranger S, Greenberg RM. 2009. Schistosoma mansoni P-glycoprotein levels increase in response to praziquantel exposure and correlate with reduced praziquantel susceptibility. Mol Biochem Parasitol 167:54–59. 165. Greenberg RM. 2013. ABC multidrug transporters in schistosomes and other parasitic flatworms. Parasitol Int 62:647–653. 166. Pica-Mattoccia L, Doenhoff MJ, Valle C, Basso A, Troiani AR, Liberti P, Festucci A, Guidi A, Cioli D. 2009. Genetic analysis of decreased praziquantel sensitivity in a laboratory strain of Schistosoma mansoni. Acta Trop 111:82–85. 167. Kasinathan RS, Morgan WM, Greenberg RM. 2010. Schistosoma mansoni express higher levels of multidrug

168.

169.

170.

171.

resistance-associated protein 1 (SmMRP1) in juvenile worms and in response to praziquantel. Mol Biochem Parasitol 173:25–31. Hines-Kay J, Cupit PM, Sanchez MC, Rosenberg GH, Hanelt B, Cunningham C. 2012. Transcriptional analysis of Schistosoma mansoni treated with praziquantel in vitro. Mol Biochem Parasitol 186:87–94. Doenhoff MJ, Hagan P, Cioli D, Southgate V, PicaMattoccia L, Botros S, Coles G, Tchuem Tchuenté LA, Mbaye A, Engels D. 2009. Praziquantel: its use in control of schistosomiasis in sub-Saharan Africa and current research needs. Parasitology 136:1825–1835. Caffrey CR, Secor WE. 2011. Schistosomiasis: from drug deployment to drug development. Curr Opin Infect Dis 24:410–417. Secor WE, Meites E, Starr MC, Workowski KA. 2014. Neglected parasitic diseases in the United States: Trichomoniasis. Am J Trop Med Hyg 90:800–804.

Susceptibility Test Methods: Parasites JACQUES LE BRAS, JÉRÔME CLAIN, AND W. EVAN SECOR

151 Accurate methods for ascertaining responses of parasites to antiparasitic drugs can prove useful at several levels. They can assist in the clinical management of individual patients, they can yield epidemiologic information that may guide drug use policies and public health interventions, and they offer crucial research tools for the development of new and better drugs. Drug susceptibility tests fall into four broad categories: in vivo tests, in vitro tests, tests with experimental animals, and molecular tests. In vivo tests with patients directly assess the clinical efficacies of existing compounds. These tests are performed in actual epidemiologic investigations, and their modest technical requirements make them suitable for use under field conditions in developing countries. The interpretation of in vivo tests is limited by potential interference by factors related to the host (e.g., immunity or variations in drug intake or metabolism) or to the environment (e.g., reinfections). However, such tests have proven instrumental in guiding drug use policies, particularly for malaria. In vitro tests circumvent these interferences by isolating the parasites from their hosts and investigating them in culture under controlled laboratory conditions, which provides opportunities for repeated assessments against multiple compounds, including experimental compounds. In vitro tests, however, are technically more demanding and therefore less amenable to performance under field conditions. They are most informative if they are used to investigate parasites that multiply rapidly in culture, a select group consisting mostly of protozoa. They are of limited use for assessing inactive drug precursors that must be activated by the host or drugs whose antiparasitic activities necessitate the synergistic effect of the host’s immune defenses. Tests with experimental animal models permit investigations of parasites that cannot be grown in culture or of drugs not yet approved for use in humans. This approach accommodates the advantages of human in vivo drug testing without the potential of exposing persons to toxic drugs that may be ineffective. However, its use is predicated on a suitable animal model for the infection and on the availability of appropriate facilities for maintaining the test animals. Furthermore, for animal models to be relevant, the pharmacokinetics of the drug under investigation should be similar in the particular model used and in humans. Molecular tests detect genetic variations that are potentially linked with resistance. Such tests offer unique advantages. PCRs can be performed with minute amounts of nonviable parasite genetic material. They can be run in

batches, allowing large-scale epidemiologic studies. Molecular analysis can circumvent potential ambiguities associated with the polyclonality of parasites infecting a single host occasionally encountered in in vivo or in vitro tests and allows the dissection of such within-host parasite populations. Because of their short duration (hours), molecular diagnostic procedures can potentially be used to guide patient management. In vivo and in vitro tests usually require more time (days to weeks) and yield results that are used mainly for epidemiologic surveillance and experimental chemotherapy studies. Drawbacks of molecular tests reside in their technical requirements and in the need to be certain that the genes evaluated correlate with functional resistance. However, thanks to the development of more-practical protocols and more-robust automated equipment, as well as a better understanding of the genes that confer resistance, molecular techniques are being used in an increasing number of laboratories, including field facilities. These different categories of tests provide complementary information. At one end of the spectrum, molecular tests analyze parasites at their most basic biologic level, without any outside interference. At the other end, in vivo tests in patients reflect complex interactions between host and parasite yet are most relevant for clinicians and public health practitioners. While a good correlation between results of various test methods is desirable, some degree of discrepancy should be expected to result from factors linked to the host or the culture conditions. Indeed, a judicious analysis of such discrepancies might provide valuable insights into the mechanisms of drug action and resistance. These points are illustrated in the following discussions of five parasitic diseases, selected for their particular chemotherapeutic challenges. A summary is provided in Table 1.

MALARIA Most drug resistance tests in malaria concern Plasmodium falciparum, the most prevalent and virulent species. Initial observations of drug-resistant malaria occur most often in a clinical context, and their confirmation is frequently sought by in vivo tests. These aim to document the parasitological and clinical response of a malaria infection in a patient treated with a standard dose of the test drug and monitored under controlled conditions. Initially standardized by the WHO for the response of P. falciparum to chloroquine, in vivo tests have been modified several times for increased performance and assessments of other drugs (1). An increasing number of

doi:10.1128/9781555817381.ch151

2563

2564 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

TABLE 1

Selected antiparasitic agents and susceptibility testing methodsa

Disease and drugs Malariab Chloroquine, amodiaquine, quinine, mefloquine, lumefantrine, piperaquine, pyronaridine, artemisinin, sulfadoxine-pyrimethamine, atovaquone-proguanil, tetracycline, primaquine, and others

Trichomoniasisc Metronidazole and tinidazole

Leishmaniasisd Sodium stibogluconate, meglumine antimoniate, pentamidine, amphotericin B, paromomycin, and miltefosine

African trypanosomiasise Pentamidine, suramin, melarsoprol, and eflornithine

Schistosomiasisf Praziquantel

Testing method(s)

Remarks

In vivo tests in patients with Plasmodium falciparum or P. vivax malaria. Culture of erythrocytic stages of P. falciparum. Criteria for assessment are (i) microscopic examination (maturation from rings to schizonts; parasite multiplication), (ii) metabolic activity (incorporation of [3H]hypoxanthine; production of pLDH, HRP2), and (iii) DNA quantitation. PCR-based genetic analysis (mostly of Plasmodium falciparum) of mutations in and amplification of genes putatively involved in resistance to chloroquine (pfcrt, pfmdr1), mefloquine (pfmdr1), antifolates (dhfr, dhps genes), atovaquone (pfcytb gene), artemisinins (kelch13 gene), and others.

Drug resistance is a major problem, especially in P. falciparum; it also occurs in P. vivax. Tests are used for epidemiologic assessment as well as for laboratory investigations. Short-term culture tests are also described for erythrocytic stages of P. vivax. On an experimental basis, there are in vitro tests to determine a drug’s effect on liver stages and sexual stages (gametocytes).

Culture under aerobic and anaerobic conditions. The criterion for assessment is parasite mobility.

Resistance to metronidazole is relative; testing is performed over a wide range of concentrations.

Culture of promastigotes. Criteria for assessment are microscopic examination with parasite counting and metabolic activity (incorporation of [3H]thymidine, hydrolysis of p-nitrophenyl phosphate, conversion of MTT or resazurin). Culture of amastigotes (intracellular or axenic). Criteria for assessment are microscopic examination with counting of stained intracellular parasites, luciferase activity of transfected parasites, flow cytometry using GFP-transfected parasites, lysis of host cells, transformation of amastigotes back to promastigotes, and promastigote assay.

Problems with most drugs are their high cost, difficulty in administration, and toxicity. There is a high level of failure of pentavalent antimonials in some areas. The choices of promastigote or amastigote assay differ with the drug being tested. Tests with intracellular or axenic amastigotes show better correlation with clinical drug efficacy but are not absolute.

Culture of trypomastigotes and assessment by microscopic examination with parasite counting, metabolic activity (hydrolysis of p-nitrophenyl phosphate, reduction of resazurin), and uptake of radiolabeled drug.

In vitro tests are used mainly for laboratory investigations. The ability to detect mutations in the transporter responsible for drug uptake may soon allow fieldapplicable tests.

Examination of damage to adult worms from experimental infections with suspected resistant strains, egg-hatching efficiency, miracidial morphology, and cercarial tail shedding.

In vivo animal tests are needed for confirmation due to the dependence of drug effect on the host immune response.

a

See the text for details. Disease caused predominantly by Plasmodium falciparum, Plasmodium vivax, Plasmodium ovale, Plasmodium malariae, and Plasmodium knowlesi. Disease caused by Trichomonas vaginalis. d Disease caused by one or several Leishmania species. e Disease caused by Trypanosoma brucei rhodesiense and T. brucei gambiense. f Disease caused predominantly by Schistosoma mansoni, Schistosoma japonicum, and Schistosoma haematobium. b c

in vivo tests comparing drug schemes have been performed since 1996, as it was necessary to assess the response of P. falciparum to artemisinin-based combined therapy (2). With regard to the fast-acting artemisinin derivatives, measurement of parasite clearance times, in addition to determination of standard clinical outcomes, has been introduced (3). Nevertheless, the diversity of study designs and analytical

methods undermines the possibility of monitoring antimalarial drug efficacy over time from diverse regions of endemicity (4). Among key variables considered to define the therapeutic response are Plasmodium species, outcome, level of immunity, and PCR adjustment to discriminate recrudescence of the initial infestation and recurrence (Plasmodium vivax) or reinfestation. Incorporating pharmacokinetic parameters is

151. Susceptibility Test Methods: Parasites n

now considered for true resistance identification within partners of artemisinins, which are frequently poorly absorbed or slowly eliminated drugs (5, 6). The standard in vitro antimalarial drug susceptibility assay determines the ex vivo growth of replicating intraerythrocytic parasites from the ring stage (the only asexual P. falciparum stage found in patient peripheral blood) to the schizont stage in 24 to 72 h in the presence of serial drug concentrations under conditions close to in vivo conditions. With regard to the standard assay, a well-suited 96-well microtiter plate format was designed using the Trager and Jensen cultivation parameters (hypoxia and buffered RPMI medium with human serum), which is the basis of the mostused tests (7, 8). The simplest test format has been adapted to field work and uses 100 μl of fingerpick capillary blood mixed with medium, a 24- to 30-h candle jar incubation, a microscopic count of multinucleated schizonts, and calculation of the 50% inhibitory concentration (IC50) and IC99 of the drug from a dose-response curve (9). Beyond this simplest format, whose reagents were prepared and commercialized through the WHO in the years 1980 to 1990, other in vitro tests that offer valuable advantages have been developed. All tests are applicable both to field-collected parasites and to the parasites growing asynchronously in long-term laboratory culture and can also be used for screening potential new antimalarials and for investigations on drug modes of action or resistance. The activities of some antimalarials, such as quinine, address only a part of the asexual erythrocytic cycle. This implies that when long-term laboratory culture of patient isolates or reference clones is used, synchronization at ring stage can improve the reproducibility of susceptibility results. Parasite growth and inhibition can be assessed using different methods. A parasite count by microscopic examination of culture smears is cumbersome and often poorly reproducible. Measurement of uptake of [3H]hypoxanthine offers a semiautomated, quantitative, high-output approach but necessitates use of radioactive material and specialized equipment in authorized laboratories, leading to high costs because of the handling of radioactive waste (10). Measurement of highly produced Plasmodium proteins, such as parasite lactate dehydrogenase (11) or histidine-rich protein 2 (12), by double-site enzymelinked immunosorbent assay demonstrated a higher sensitivity than use of radioisotopes, although these tests are timeconsuming and commercial kits are costly (13). Other tests measure the production of DNA during the maturation of parasites using SYBR green I fluorescent dye (14–16). The successful completion and interpretation of all in vitro tests depends on several factors that in turn depend on samples, materials, culture, and statistical methods to generate inhibitory constants. Sample factors to be considered include a recent intake of antimalarial drugs by the patient, which may decrease the test success rate; high parasite inocula, which can lead to an overestimation of resistance to some drugs (17); and folate and para-aminobenzoic acid in the culture medium, which antagonize the in vitro effect of antifolate drugs (18). Another sample factor is the short life of erythrocyte P. falciparum parasites outside the host; each day that passes at 4°C halves their capacity to survive and grow in vitro (19). Processes include the necessity of preparing and distributing dilutions of drugs in wells of plates, which results in particular difficulties for drugs other than chloroquine, as most are poorly soluble or have limited shelf lives. Culture necessitates supplementation of RPMI medium with human serum or AlbuMAX, although this can affect IC50s, as they interact with drugs differently (20). All these fundamental methodological issues undermine ac-

2565

curate comparisons of in vitro susceptibilities either between laboratories or within a single laboratory over time. Consensus exists on parameters of culture, and suggested improvements include measures of quality control of key parameters, such as predosed plates (titrated drug solutions) and reference of endpoints of isolate susceptibilities to those of reference clones with known susceptibilities (21). Finally, standardized mathematical analysis of concentration inhibition assays is possible through free Web-based tools (22, 23). Drug concentrations associated with the IC50 are determined by a modified sigmoid maximum-effect (Emax) modelfitting algorithm and display the precision of IC50 estimation. The standard in vitro antimalarial drug susceptibility assay described above, however, does not permit accurate potency measurement of the fast-acting and stage-specific artemisinin derivatives for which specific protocols have been developed recently (24–26). These new assays test the capacity of early-ring-stage parasites to survive a brief pulse exposure to high artemisinin concentrations (the so-called ring-stage survival assay [RSA]), better mimicking the situation faced by parasites in vivo. In a study with Cambodian parasite isolates, the survival rate estimated with the RSA correlates with in vivo parasite clearance half-lives (25). Tests exploring antimalarial efficacy at various stages of the Plasmodium life cycle have been designed but are not adapted for routine laboratory use (27). However, the microtechnique has been adapted for testing of erythrocytic stages of P. vivax (28), and cultures of the liver and sexual blood stages of P. falciparum can also be used, although they are substantially more cumbersome (29, 30). Genetic markers that now offer tools for assessing parasite drug resistance have been identified for several major antimalarials. Chloroquine resistance has been linked to a P. falciparum chloroquine resistance transporter (PfCRT) K76T change on a transmembrane channel in the digestive vacuole of P. falciparum, following a mutation of the pfcrt gene (31, 32). P. falciparum multidrug resistance gene 1 (pfmdr1) encodes a P glycoprotein homologue of a human ABC transporter that transports toxic compounds across the digestive vacuole membrane. Point mutations and gene amplification of pfmdr1 have been linked, to various extents, to altered susceptibilities to various antimalarials (chloroquine and other 4-aminoquinolines, quinine and other amino-alcohols, and artemisinin derivatives) (33–35). Regarding the artemisinins, missense mutations in the kelch13 gene have been proposed as in vitro and in vivo markers of artemisinin resistance (36). Resistance to antifolates, such as sulfadoxine-pyrimethamine, has been associated with S108N, C59R, and N51I changes in P. falciparum dihydrofolate reductase (37–39) or its homologues in P. vivax (40). High levels of resistance to antifolate drugs are associated with the P. falciparum dihydrofolate reductase I164L change (37, 41) or changes in the P. falciparum dihydropteroate synthase target of sulfa drugs (42). Resistance to atovaquone-proguanil has been linked to substitutions in codon 268 of the P. falciparum cytochrome b (pfcytb) gene, resulting in the amino acid change S, N, or C, leading to highlevel resistance (43). Such genetic polymorphisms can be identified in individual parasite isolates by various standard techniques, which include mutation-specific nested PCR (44) or PCR followed by sequencing (45), restriction fragment length polymorphism analysis (46), and single-nucleotide primer extension with detection of fluorescent products on a capillary sequencer (47), or by using a DNA microarraybased method (48). Next-generation sequencing also allows accurate detection of drug resistance genotypes from pooled parasite isolates (49). Possibilities for analyzing haplotypes

2566 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

in mixed parasite isolates may improve our ability to find the clinical relevance of combined mutations (50). As long as validated molecular markers are lacking for resistance to artemisinin’s partner drugs, in vivo and in vitro surveillance will remain critical surveillance tools for the emergence of resistance to artemisinin-based combined therapy (51, 52). The judicious use of in vivo, in vitro, and molecular tests can yield valuable information that will help us to adapt drug policies before extension of resistance brings about elevated morbidity consequences (53, 54).

TRICHOMONIASIS Trichomonas vaginalis is the most common nonviral sexually transmitted infection in the United States and throughout the world; however, metronidazole and tinidazole are the only drugs currently approved by the U.S. Food and Drug Administration to treat trichomoniasis. With 4.3% of T. vaginalis-infected sexually transmitted disease clinic patients in the United States demonstrating some degree of resistance to metronidazole, standard treatment regimens may not be effective for approximately 159,000 individuals (55, 56). Fortunately, drug susceptibility testing has proven useful in identifying alternative treatment protocols that are usually successful in effecting patient cure (57). In vitro testing for resistance to 5-nitroimidazoles is indicated following failure of two standard treatments to cure the patient’s infection. Susceptibility testing for T. vaginalis is a simple assay of parasite motility in the presence of drug (58). Axenic trichomonads are cultured in Diamond’s Trypticase-yeastmaltose medium with serial dilutions (400 to 0.2 μg/ml) of metronidazole or tinidazole dissolved in dimethyl sulfoxide (DMSO) and the appropriate parallel concentrations of DMSO in U-bottom microtiter plates. Plates are incubated at 37°C for 48 h and are then examined microscopically with an inverted phase-contrast microscope. The lowest concentration of drug in which no motile organisms are observed is defined as the minimum lethal concentration. Minimal lethal concentrations greater than 100 μg of metronidazole per ml have been associated with clinical resistance (59). This assay has been adapted to determine IC50s and screen novel compounds for antitrichomonad activity by measuring incorporation of [3H]thymidine (60, 61) or acid phosphatase activity (62). These modifications have not as yet been adapted to monitor clinically resistant isolates, but theoretically, this could be accomplished rather easily. One difficulty of the in vitro culture method for assessing the resistance of T. vaginalis isolates is the need to derive axenic cultures from clinical specimens. As individuals with trichomoniasis are often infected with other sexually transmitted disease organisms that also grow in Diamond’s medium, this may require an extended time. Molecular comparisons of T. vaginalis isolates suggest that genetic markers for metronidazole resistance could be identified (63–66). The prospect of a molecular marker for resistant T. vaginalis is exciting, as it may make possible a PCR-based assay for resistance. This would obviate the need for establishing axenic cultures and may allow direct testing of specimens from patients suspected of harboring a resistant isolate.

LEISHMANIASIS Treatment failure in leishmaniasis may result from either true drug resistance or patient factors, such as poor drug absorption, treatment compliance, or immunodeficiencies,

that preclude effective chemotherapeutic action. Differentiation of these two possibilities is important for both individual patient care and general public health. Traditional testing for drug resistance in Leishmania is performed by culturing the promastigote form of the parasite with drug in standard cell culture media (Schneider’s medium, RPMI medium, or M199 medium) supplemented with bovine sera for 42 to 72 h at 26 to 37°C and by assessing viability by direct counting (67), [3H]thymidine incorporation (68), enzymatic hydrolysis of p-nitrophenyl phosphate (69), conversion of 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT) (70), or reduction of resazurin (71). [3H]thymidine incorporation is perhaps the most sensitive method, but it requires access to radioactive materials and a scintillation counter. Hydrolysis of p-nitrophenyl phosphate and conversion of MTT are colorimetric assays, but p-nitrophenyl phosphate hydrolysis has high backgrounds caused by medium components and MTT may interact with certain drugs (e.g., meglumine antimoniate). An advantage of resazurin (also known as Alamar Blue) is that it is not necessary to further manipulate the parasites in order to read the assay. Thus, drug activities can be monitored at different time points after initiation of cultures. Although promastigote assays are the easiest to use, they yield clinically reliable results only for drugs (such as miltefosine and amphotericin B) that do not require cellular mechanisms for activation (72, 73). In contrast, the more widely used pentavalent antimonials require reduction by host cells to the active trivalent form. As a result, promastigote assays correlate poorly with clinical resistance, and assays that test drug susceptibility of the intracellular amastigote form of leishmania parasites are recommended (74, 75). Unfortunately, intracellular assays are very labor-intensive. Macrophages are infected with promastigotes that must then transform into amastigotes prior to exposure to drug. Each test condition necessitates preparation of slides, which are then stained for microscopic counting of the proportion of infected macrophages and the number of amastigotes per macrophage in drug-treated and control cultures. Furthermore, promastigotes from field isolates are not uniformly infective for macrophages, leading to selection bias that may reduce the correlation between drug susceptibility assays and clinical treatment outcome (76, 77). Intracellular assays have been facilitated by stable transformation of isolates with firefly luciferase. Cells are simply lysed, the substrate luciferin is added, and activity is measured on a luminometer (78). Similarly, transfection of isolates with the gene encoding green fluorescent protein (GFP) can be used (79). Following incubation with the drug, parasites are enumerated on a flow cytometer, with no need for addition of substrates because GFP is intrinsically fluorescent. An advantage of transfection assays is the ability to compare drug activities against promastigotes, axenic amastigotes, and intracellular amastigotes. However, these assays are more useful for screening experimental compounds rather than testing field isolates because of the challenges of standardized transfection between isolates and the expensive equipment needed to measure the results. If transfection with exogenous genes is undesirable, mammalian cells that have been infected with parasites and treated with drug can be fixed and permeabilized. Intracellular amastigotes are then detected with anti-Leishmania lipophosphoglycan monoclonal antibody followed by a fluorescent-marker-labeled anti-mouse immunoglobulin antibody (80). This assay can be used for assessing the drug sensitivities of a variety of Leishmania species, and it is adaptable to use with individual patient isolates. Alternatively, following

151. Susceptibility Test Methods: Parasites n

incubation of infected cells with a drug, controlled lysis of macrophages can be performed to release amastigotes that are then transformed back into promastigotes and enumerated using one of the microscopic, [3H]thymidine incorporation, colorimetric, or fluorometric assays described above (81). As molecular mechanisms of drug resistance in Leishmania spp. become better defined, it may be possible to utilize PCR and DNA sequencing of single nucleotide polymorphisms (SNPs) for epidemiologic studies of resistant isolates. Resistance-associated SNPs from both visceral and cutaneous Leishmania species have been identified in genes for drug transporters, the stress response, and thiol and redox metabolism (82, 83). These assays are of interest, as no manipulation beyond growing parasites is needed. However, it is likely that applicability will be species and drug restricted.

AFRICAN TRYPANOSOMIASIS As in leishmania drug sensitivity testing, trypanosomes can be cultured in vitro with drug and monitored for viability by enzymatic hydrolysis of p-nitrophenyl phosphate (69) or by direct counting (84). Parasites are cultured at 37°C for 24 h in 4 to 5% CO2 in phenol red-free Iscove’s medium containing hypoxanthine, thymidine, glutamine, L-cysteine, pyruvate, β-mercaptoethanol, and heat-inactivated bovine or horse sera. An in vitro lysis assay has been developed for testing the sensitivities of Trypanosoma brucei subspecies to melarsoprol (85). It requires the ability to culture trypanosomes and a thermostatically controlled recording spectrophotometer. As susceptible organisms die, their absorbance at 500 nm is reduced over the course of 30 min. If drugsusceptible and drug-resistant strains are available to use as controls, this technique may perhaps be adapted to field use because differences in intensity at this wavelength can be distinguished by the unaided eye. Visible color change can also be observed when trypanosomes are incubated with resazurin (86). However, unlike in assays for Leishmania drug sensitivity, the dye is added after parasites are incubated with drugs for 66 to 72 h, depending on the parasite subspecies. Because drug resistance in African trypanosomes is primarily a function of reduced drug uptake, radiolabeled drugs can be incubated with parasites to identify resistant isolates, with reduced cellular incorporation of radioactivity suggesting resistance (87). Similarly, because drug uptake is reduced in isolates with point mutations or deletions of aminopurine transporters, it may soon be possible to utilize molecular tools to identify trypanosome isolates that are drug resistant (88). One of the genetic mutations leading to an amino acid change in the transporter responsible for melarsoprol uptake abrogates a restriction endonuclease site, while a different mutation creates one (89). Thus, PCR amplification of the appropriate gene followed by enzymatic digestion and gel electrophoresis results in different DNA banding patterns that might be used to distinguish melarsoprol-susceptible and -resistant isolates. While more clinical drugresistant isolates must be evaluated to confirm the utility of this test, a similar banding pattern between melarsoprolresistant laboratory-derived and field isolates (89) suggests that this tool may be useful for both management of individual patients’ infections and tracking the spread of resistance in a population (90).

SCHISTOSOMIASIS Drug resistance testing of schistosomes differs greatly from that of protozoan parasites in both purpose and methods.

2567

Adult worms do not replicate within the definitive host, thus circumventing one of the mechanisms associated with rapid development of drug resistance. As a result, widespread resistance to praziquantel, the drug of choice for schistosomiasis, has not emerged even under heavy drug pressure (91– 93). Thus, the need to evaluate drug resistance for individual infections is rare. However, because reliance on a sole drug is risky and schistosomes have developed resistance to other drugs, techniques to monitor praziquantel resistance are needed. The failure of adult worms to replicate makes drug susceptibility testing methods more challenging. Investigation of drug resistance has typically been performed by (i) obtaining eggs from an unsuccessfully treated mammalian host, (ii) infecting the appropriate intermediate snail host, (iii) isolating cercariae from the infected snail, and (iv) infecting and treating experimental animals. A protocol to determine the 50% effective dose in mice has been developed and demonstrates good reproducibility among different laboratories (94). This approach has the drawbacks of being very time-consuming and technically challenging, especially in areas of endemicity that may not have well-developed experimental snail and rodent colonies. Additionally, resistance phenotypes may not persist in the absence of drug pressure, clouding our ability to interpret results (95). Fiftypercent effective doses can also be estimated in an in vitro assay by assessing contraction of worms following perfusion and incubation with praziquantel and correspond well with in vivo results (96, 97). Although this approach is less complicated than infecting, treating, and perfusing additional mice, it still requires the availability of naive snails and some mice to obtain the adult worms, along with the time and expertise needed to perpetuate the life cycle. In addition, the parasites that successfully infect mice may not accurately represent the diversity of field isolates. Recently, an in vitro method that circumvents these problems has been adapted to field use (98, 99). Eggs from stools of infected individuals are isolated and hatched to release miracidia, which are then exposed to praziquantel, and effects on morphology are monitored. The miracidia from persons who then cleared their infection following treatment were more affected by the drug than the miracidia from stools whose donors did not clear their infections. Evaluating changes in the parasite’s population structure using miracidia obtained from eggs in patients’ stools is another approach to monitor development of potential drug resistance. Changes in the genetic composition of schistosomes following treatment suggest praziquantel efficacy, while maintenance of the same microsatellite distribution may indicate treatment failure (100, 101). Ongoing surveillance for development of drug resistance will be an important component for schistosomiasis control programs that are dependent on mass drug administration.

FUTURE PERSPECTIVES Most susceptibility tests for antiparasitic drugs are not routinely available in clinical diagnostic laboratories, because these procedures are not in frequent demand and present special technical requirements. Such tests are performed mainly in reference or research laboratories or during epidemiologic investigations in areas of endemicity. Thanks to recent advances in laboratory technology and genetic analysis of parasites, available tests are increasingly sophisticated and informative. The development of tests that are robust and simple to use will facilitate field studies that aim at optimizing the deployment of currently available drugs.

2568 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

These field tests usefully complement the more sophisticated procedures used in research laboratories whose main orientation is toward the development of novel antiparasitic compounds and deciphering drug resistance mechanisms.

REFERENCES 1. World Health Organization. 2003. Assessment and Monitoring of Antimalarial Drug Efficacy for the Treatment of Uncomplicated Falciparum Malaria. WHO/HTM/RBM/ 2003.50. World Health Organization, Geneva, Switzerland. 2. World Health Organization. 2005. Susceptibility of P. falciparum to Antimalarial Drugs: Report on Global Monitoring: 1996–2004. WHO/HTM/MAL/2005.1103. World Health Organization, Geneva, Switzerland. 3. Flegg JA, Guerin PJ, White NJ, Stepniewska K. 2011. Standardizing the measurement of parasite clearance in falciparum malaria: the parasite clearance estimator. Malar J 10:339. 4. Price RN, Dorsey G, Ashley EA, Barnes KI, Baird JK, d’Alessandro U, Guerin PJ, Laufer MK, Naidoo I, Nosten F, Olliaro P, Plowe CV, Ringwald P, Sibley CH, Stepniewska K, White NJ. 2007. World Antimalarial Resistance Network. I: Clinical efficacy of antimalarial drugs. Malar J 6:119. 5. White NJ. 2002. The assessment of antimalarial drug efficacy. Trends Parasitol 18:458–464. 6. Barnes KI, Watkins WM, White NJ. 2008. Antimalarial dosing regimens and drug resistance. Trends Parasitol 24: 127–134. 7. Trager W, Jensen JB. 1976. Human malaria parasites in continuous culture. Science 193:673–675. 8. Rieckmann KH, Sax LJ, Campbell GH, Mrema JE. 1978. Drug sensitivity of Plasmodium falciparum. An in-vitro microtechnique. Lancet i:22–23. 9. World Health Organization. 2001. In Vitro Micro-Test (Mark III) for the Assessment of P. falciparum Susceptibility to Chloroquine, Mefloquine, Quinine, Amodiaquine, Sulfadoxine/Pyrimethamine and Artemisinin. CTD/MAL/9720 Rev 2. World Health Organization, Geneva, Switzerland. 10. Desjardins RE, Canfield CJ, Haynes JD, Chulay JD. 1979. Quantitative assessment of antimalarial activity in vitro by a semiautomated microdilution technique. Antimicrob Agents Chemother 16:710–718. 11. Moreno A, Brasseur P, Cuzin-Ouattara N, Blanc C, Druilhe P. 2001. Evaluation under field conditions of the colourimetric DELI-microtest for the assessment of Plasmodium falciparum drug resistance. Trans R Soc Trop Med Hyg 95:100–103. 12. Noedl H, Attlmayr B, Wernsdorfer WH, Kollaritsch H, Miller RS. 2004. A histidine-rich protein 2-based malaria drug sensitivity assay for field use. Am J Trop Med Hyg 71:711–714. 13. Kaddouri H, Nakache S, Houze S, Mentre F, Le Bras J. 2006. Susceptibility of Plasmodium falciparum clinical isolates from Africa using a highly sensitive Plasmodium lactate dehydrogenase immunodetection assay and inhibitory maximum effect model for precise measurement of the 50-percent inhibitory concentration. Antimicrob Agents Chemother 50:3343–3349. 14. Smilkstein M, Sriwilaijaroen N, Kelly JX, Wilairat P, Riscoe M. 2004. Simple and inexpensive fluorescencebased technique for high-throughput antimalarial drug screening. Antimicrob Agents Chemother 48:1803–1806. 15. Bacon DJ, Latour C, Lucas C, Colina O, Ringwald P, Picot S. 2007. Comparison of a SYBR green I-based assay with a histidine-rich protein II enzyme-linked immunosorbent assay for in vitro antimalarial drug efficacy testing and application to clinical isolates. Antimicrob Agents Chemother 51:1172–1178.

16. Rason MA, Randriantsoa T, Andrianantenaina H, Ratsimbasoa A, Ménard D. 2008. Performance and reliability of the SYBR Green I based assay for the routine monitoring of susceptibility of P. falciparum clinical isolates. Trans R Soc Trop Med Hyg 102:346–351. 17. Duraisingh MT, Jones P, Sambou I, von Seidlein L, Pinder M, Warhurst DC. 1999. Inoculum effect leads to overestimation of in vitro resistance for artemisinin derivatives and standard antimalarials: a Gambian field study. Parasitology 119:435–440. 18. Wang P, Sims PF, Hyde JE. 1997. A modified in vitro sulfadoxine susceptibility assay for Plasmodium falciparum suitable for investigating Fansidar resistance. Parasitology 115:223–230. 19. Basco LK. 2004. Molecular epidemiology of malaria in Cameroon. XX. Experimental studies on various factors of in vitro drug sensitivity assays using fresh isolates of P. falciparum. Am J Trop Med Hyg 70:474–480. 20. Basco LK. 2003. Molecular epidemiology of malaria in Cameroon. XV. Experimental studies on serum substitutes and supplements and alternative culture media for in vitro drug sensitivity assays using fresh isolates of P. falciparum. Am J Trop Med Hyg 69:168–173. 21. Bacon DJ, Jambou R, Fandeur T, Le Bras J, Wongsrichanalai C, Fukuda MM, Ringwald P, Sibley CH, Kyle DE. 2007. An in vitro antimalarial drug susceptibility database: a component of the World Antimalarial Resistance Network (WARN) database. Malar J 6:120–124. 22. Le Nagard H, Vincent C, Mentré F, Le Bras J. 2011. Online analysis of in vitro resistance to antimalarial drugs through nonlinear regression. Comput Methods Programs Biomed 104:10–18. 23. Woodrow CJ, Dahlström S, Cooksey R, Flegg JA, Le Nagard H, Mentré F, Murillo C, Ménard D, Nosten F, Sriprawat K, Musset L, Quashie NB, Lim P, Fairhurst RM, Nsobya SL, Sinou V, Noedl H, Pradines B, Johnson JD, Guerin PJ, Sibley CH, Le Bras J. 2013. High-throughput analysis of antimalarial susceptibility data by the WorldWide Antimalarial Resistance Network (WWARN) in vitro analysis and reporting tool. Antimicrob Agents Chemother 57:3121–3130. 24. Klonis N, Xie SC, McCaw JM, Crespo-Ortiz MP, Zaloumis SG, Simpson JA, Tilley L. 2013. Altered temporal response of malaria parasites determines differential sensitivity to artemisinin. Proc Natl Acad Sci USA 110:5157– 5162. 25. Witkowski B, Amaratunga C, Khim N, Sreng S, Chim P, Kim S, Lim P, Mao S, Sopha C, Sam B, Anderson JM, Duong S, Chuor CM, Taylor WR, Suon S, MercereauPuijalon O, Fairhurst RM, Ménard D. 2013. Novel phenotypic assays for the detection of artemisinin-resistant Plasmodium falciparum malaria in Cambodia: in-vitro and ex-vivo drug response studies. Lancet Infect Dis 13:1043– 1049. 26. Witkowski B, Khim N, Chim P, Kim S, Ke S, Kloeung N, Chy S, Duong S, Leang R, Ringwald P, Dondorp AM, Tripura R, Benoit-Vical F, Berry A, Gorgette O, Ariey F, Barale JC, Mercereau-Puijalon O, Ménard D. 2013. Reduced artemisinin susceptibility of Plasmodium falciparum ring stages in western Cambodia. Antimicrob Agents Chemother 57:914–923. 27. Delves M, Plouffe D, Scheurer C, Meister S, Wittlin S, Winzeler EA, Sinden RE, Leroy D. 2012. The activities of current antimalarial drugs on the life cycle stages of Plasmodium: a comparative study with human and rodent parasites. PLoS Med 9:e1001169. doi:10.1371/journal. pmed.1001169. 28. Tasanor O, Noedl H, Na-Banchang K, Congpuong K, Sirichaisinthop J, Wernsdorfer WH. 2002. An in vitro system for assessing the sensitivity of Plasmodium vivax to chloroquine. Acta Trop 83:49–61.

151. Susceptibility Test Methods: Parasites n 29. Stahel E, Mazier D, Guillouzo A, Miltgen F, Landau I, Mellouk S, Beaudoin RL, Langlois P, Gentilini M. 1988. Iron chelators: in vitro inhibitory effect on the liver stage of rodent and human malaria. Am J Trop Med Hyg 39:236– 240. 30. Coleman RE, Nath AK, Schneider I, Song G-H, Klein TA, Milhous WK. 1994. Prevention of sporogony of Plasmodium falciparum and P. berghei in Anopheles stephensi mosquitoes by transmission-blocking antimalarials. Am J Trop Med Hyg 50:646–653. 31. Fidock DA, Nomura T, Talley AK, Cooper RA, Dzekunov SM, Ferdig MT, Ursos LMB, Sidhu ABS, Naudé B, Deitsch KW, Su X, Wootton JC, Roepe PD, Wellems TE. 2000. Mutations in the P. falciparum digestive vacuole transmembrane protein PfCRT and evidence for their role in chloroquine resistance. Mol Cell 6:861–871. 32. Ecker A, Lehane AM, Clain J, Fidock DA. 2012. PfCRT and its role in antimalarial drug resistance. Trends Parasitol 28:504–511. 33. Pickard AL, Wongsrichanalai C, Purfield A, Kamwendo D, Emery K, Zalewski C, Kawamoto F, Miller RS, Meshnick SR. 2003. Resistance to antimalarials in Southeast Asia and genetic polymorphisms in pfmdr1. Antimicrob Agents Chemother 47:2418–2423. 34. Price RN, Uhlemann AC, Brockman A, McGready R, Ashley E, Phaipun L, Patel R, Laing K, Looareesuwan S, White NJ, Nosten F, Krishna S. 2004. Mefloquine resistance in Plasmodium falciparum and increased pfmdr1 gene copy number. Lancet 364:438–447. 35. Valderramos SG, Fidock DA. 2006. Transporters involved in resistance to antimalarial drugs. Trends Pharmacol Sci 27:594–601. 36. Ariey F, Witkowski B, Amaratunga C, Beghain J, Langlois AC, Khim N, Kim S, Duru V, Bouchier C, Ma L, Lim P, Leang R, Duong S, Sreng S, Suon S, Chuor CM, Bout DM, Ménard S, Rogers WO, Genton B, Fandeur T, Miotto O, Ringwald P, Le Bras J, Berry A, Barale JC, Fairhurst RM, Benoit-Vical F, Mercereau-Puijalon O, Ménard D. 2014. A molecular marker of artemisininresistant Plasmodium falciparum malaria. Nature 505:50– 55. 37. Peterson DS, Walliker D, Wellems TE. 1988. Evidence that a point mutation in dihydrofolate reductase-thymidylate synthase confers resistance to pyrimethamine in falciparum malaria. Proc Natl Acad Sci USA 85:9114–9118. 38. Cowman AF, Morry MJ, Biggs BA, Cross GA, Foote SJ. 1988. Amino acid changes linked to pyrimethamine resistance in the dihydrofolate reductase-thymidylate synthase gene of Plasmodium falciparum. Proc Natl Acad Sci USA 85:9109–9113. 39. Müller IB, Hyde JE. 2013. Folate metabolism in human malaria parasites—75 years on. Mol Biochem Parasitol 188:63–77. 40. Hastings MD, Porter KM, Maguire JD, Susanti I, Kania W, Bangs MJ, Sibley CH, Baird JK. 2003. Dihydrofolate reductase mutations in Plasmodium vivax from Indonesia and therapeutic response to sulfadoxine plus pyrimethamine. J Infect Dis 189:744–750. 41. Sirawaraporn W, Sathitkul T, Sirawaraporn R, Yuthavong Y, Santi DV. 1997. Antifolate-resistant mutants of Plasmodium falciparum dihydrofolate reductase. Proc Natl Acad Sci USA 94:1124–1129. 42. Nzila AM, Mberu EK, Sulo J, Dayo H, Winstanley PA, Sibley CH, Watkins WM. 2000. Towards an understanding of the mechanism of pyrimethamine-sulfadoxine resistance in Plasmodium falciparum: genotyping of dihydrofolate reductase and dihydropteroate synthase of Kenyan parasites. Antimicrob Agents Chemother 44:991–996. 43. Musset L, Bouchaud O, Matheron S, Massias L, Le Bras J. 2006. Clinical atovaquone-proguanil resistance of Plas-

44.

45.

46.

47.

48.

49.

50.

51.

52.

53.

54.

55.

56.

57.

2569

modium falciparum associated with cytochrome b codon 268 mutations. Microbes Infect 9:2599–2604. Djimdé A, Doumbo OK, Cortese JF, Kayentao K, Doumbo S, Diourté Y, Dicko A, Su X, Nomura T, Fidock DA, Wellems TE, Plowe CV. 2001. A molecular marker for chloroquine-resistant falciparum malaria. N Engl J Med 344:257–263. Basco LK, Ringwald P. 2000. Molecular epidemiology of malaria in Yaounde, Cameroon. VI. Sequence variations in the Plasmodium falciparum dihydrofolate reductase-thymidylate synthase gene and in vitro resistance to pyrimethamine and cycloguanil. Am J Trop Med Hyg 62:271–276. Syafruddin D, Asih PB, Casey GJ, Maguire J, Baird JK, Nagesha HS, Cowman AF, Reeder JC. 2005. Molecular epidemiology of Plasmodium falciparum resistance to antimalarial drugs in Indonesia. Am J Trop Med Hyg 72:174– 181. Nair S, Brockman A, Paiphun L, Nosten F, Anderson TJ. 2002. Rapid genotyping of loci involved in antifolate drug resistance in Plasmodium falciparum by primer extension. Int J Parasitol 32:852–858. Crameri A, Marfurt J, Mugittu K, Maire N, Regös A, Coppee JY, Sismeiro O, Burki R, Huber E, Laubscher D, Puijalon O, Genton B, Felger I, Beck HP. 2007. Rapid microarray-based method for monitoring of all currently known single-nucleotide polymorphisms associated with parasite resistance to antimalaria drugs. J Clin Microbiol 45:3685–3691. Taylor SM, Parobek CM, Aragam N, Ngasala BE, Mårtensson A, Meshnick SR, Juliano JJ. 2013. Pooled deep sequencing of Plasmodium falciparum isolates: an efficient and scalable tool to quantify prevailing malaria drugresistance genotypes. J Infect Dis 208:1998–2006. Hastings IM, Smith TA. 2008. MalHaploFreq: a computer programme for estimating malaria haplotype frequencies from blood samples. Malar J 7:130. Dondorp AM, Nosten F, Yi P, Das D, Phyo AP, Tarning J, Lwin KM, Ariey F, Hanpithakpong W, Lee SJ, Ringwald P, Silamut K, Imwong M, Chotivanich K, Lim P, Herdman T, An SS, Yeung S, Singhasivanon P, Day NP, Lindegardh N, Socheat D, White NJ. 2009. Artemisinin resistance in Plasmodium falciparum malaria. N Engl J Med 361:455–467. Lim P, Dek D, Try V, Eastman RT, Chy S, Sreng S, Suon S, Mao S, Sopha C, Sam B, Ashley EA, Miotto O, Dondorp AM, White NJ, Su XZ, Char MC, Anderson JM, Amaratunga C, Menard D, Fairhurst RM. 2013. Ex vivo susceptibility of Plasmodium falciparum to antimalarial drugs in western, northern, and eastern Cambodia, 2011– 2012: association with molecular markers. Antimicrob Agents Chemother 57:5277–5283. Sibley C, Barnes KI, Plowe CV. 2007. The rationale for creating a world antimalarial resistance network. Malar J 6:1–3. Vestergaard LS, Ringwald P. 2007. Responding to the challenge of antimalarial drug resistance by routine monitoring to update national malaria treatment policies. Am J Trop Med Hyg 77:153–159. Kirkcaldy RD, Augostini P, Asbel LE, Bernstein KT, Kerani RP, Mettenbrink CJ, Pathela P, Schwebke JR, Secor WE, Workowski KA, Davis D, Braxton J, Weinstock HS. 2012. Trichomonas vaginalis antimicrobial resistance in 6 US cities, STD Surveillance Network, 2009– 2010. Emerg Infect Dis 18:939–943. Secor WE. 2012. Trichomonas vaginalis: treatment questions and challenges. Expert Rev Anti Infect Ther 10:107– 109. Bosserman EA, Helms DJ, Mosure DJ, Secor WE, Workowski KA. 2011. Utility of antimicrobial susceptibility testing in Trichomonas vaginalis infected women with clinical treatment failure. Sex Transm Dis 38:983–987.

2570 n

ANTIPARASITIC AGENTS AND SUSCEPTIBILITY TEST METHODS

58. Meingassner JG, Thurner J. 1979. Strain of Trichomonas vaginalis resistant to metronidazole and other 5-nitroimidazoles. Antimicrob Agents Chemother 15:254–257. 59. Müller M, Lossick JG, Gorrell TE. 1988. In vitro susceptibility of Trichomonas vaginalis and treatment outcome in vaginal trichomoniasis. Sex Transm Dis 15:17–24. 60. Crowell AL, Stephens CE, Kumar A, Boykin DE, Secor WE. 2004. Evaluation of dicationic compounds for activity against Trichomonas vaginalis.Antimicrob Agents Chemother 48:3602–3605. 61. Goodhew EB, Secor WE. 2013. Drug library screening against metronidazole-sensitive and -resistant Trichomonas vaginalis isolates. Sex Transm Infect 89:479–484. 62. Martínez-Grueiro MM, Montero-Pereira D, GiménezPardo C, Nogal-Ruiz JJ, Escario JA, Gómez-Barrio A. 2003. Trichomonas vaginalis: determination of acid phosphatase activity as a pharmacological screening procedure. J Parasitol 89:1076–1077. 63. Hampl V, Vanˇ ácˇ ová S, Kulda J, Fleger J. 2001. Concordance between genetic relatedness and phenotypic similarities of Trichomonas vaginalis strains. BMC Evol Biol 1:11. 64. Snipes LJ, Gamard PM, Narcisi EM, Beard CB, Lehmann T, Secor WE. 2000. Molecular epidemiology of metronidazole resistance in a population of Trichomonas vaginalis clinical isolates. J Clin Microbiol 38:3004–3009. 65. Meade JC, de Mestral J, Stiles JK, Secor WE, Finley RW, Cleary JD, Lushbaugh WB. 2009. Genetic diversity of Trichomonas vaginalis clinical isolates determined by EcoR1 restriction fragment length polymorphism of heat-shock protein 70 genes. Am J Trop Med Hyg 80:245–251. 66. Conrad M, Gorman A, Schillinger JA, Fiori PL, Arroyo R, Malla N, Dubey ML, Gonzalez J, Blank S, Secor WE, Carlton JM. 2012. Population genetics of the sexually transmitted pathogen Trichomonas vaginalis and evidence for sexual recombination. PLoS Negl Trop Dis 6:e1573. doi:10.1371/journal.pntd.0001573. 67. Callahan HL, Portal AC, Devereaux R, Grögl M. 1997. An axenic amastigote system for drug screening. Antimicrob Agents Chemother 41:818–822. 68. Grögl M, Thomason TN, Franke ED. 1992. Drug resistance in leishmaniasis: its implication in systemic chemotherapy of cutaneous and mucocutaneous disease. Am J Trop Med Hyg 47:117–126. 69. Bodley AL, McGarry MW, Shapiro TA. 1995. Drug cytotoxicity assay for African trypanosomes and Leishmania species. J Infect Dis 172:1157–1159. 70. Sereno D, Lemesre J-L. 1997. Axenically cultured amastigote forms as an in vitro model for investigation of antileishmanial agents. Antimicrob Agents Chemother 41:972– 976. 71. Mikus J, Steverding D. 2000. A simple colorimetric method to screen drug cytotoxicity against Leishmania using the dye Alamar Blue®. Parasitol Int 48:265–269. 72. Vermeersch M, da Luz RI, Toté K, Timmermans J-P, Cos P, Maes L. 2009. In vitro susceptibilities of Leishmania donovani promastigote and amastigote stages to antileishmanial reference drugs: practical relevance to stage-specific differences. Antimicrob Agents Chemother 53:3855–3859. 73. Kulshrestha A, Bhandari V, Mukhopadhyay R, Ramesh V, Sundar S, Maes L, Dujardin JC, Roy S, Salotra P. 2013. Validation of a simple-resazurin-based promastigote assay for the routine monitoring of miltefosine susceptibility in clinical isolates of Leishmania donovani. Parasitol Res 112:825–828. 74. Lira R, Sundar S, Makharia A, Kenney R, Gam A, Saraiva E, Sacks D. 1999. Evidence that the high incidence of treatment failures in Indian kala-azar is due to the emergence of antimony-resistant strains of Leishmania donovani. J Infect Dis 180:564–567. 75. Aït-Oudhia K, Gazanion E, Vergnes B, Oury B, Sereno D. 2011. Leishmania antimony resistance: what we know

76.

77.

78.

79.

80.

81.

82.

83.

84.

85.

86.

87.

88.

89.

90.

what we can learn from the field. Parasitol Res 109:1225– 1232. da Luz RI, Vermeersch M, Dujardin J-C, Cos P, Maes L. 2009. In vitro sensitivity testing of Leishmania field isolates: preconditioning of promastigotes enhances infectivity for macrophage host cells. Antimicrob Agents Chemother 53: 5197–5203. Rijal S, Yardley V, Chappuis F, Decuypere S, Khanal B, Singh R, Boelaert M, De Doncker S, Croft S, Dujardin J-C. 2007. Antimonial treatment of visceral leishmaniasis: are current in vitro susceptibility assays adequate for prognosis of in vivo therapy outcome? Microb Infect 9:529– 535. Sereno D, Roy G, Lemesre JL, Papadopoulou B, Ouellette M. 2001. DNA transformation of Leishmania infantum axenic amastigotes and their use in drug screening. Antimicrob Agents Chemother 45:1168–1173. Singh N, Dube A. 2004. Fluorescent Leishmania: application to anti-leishmanial drug testing. Am J Trop Med Hyg 71:400–402. Di Giorgio C, Ridoux O, Delmas F, Azas N, Gasquet M, Timon-David P. 2000. Flow cytometric detection of Leishmania parasites in human monocyte-derived macrophages: application to antileishmanial-drug testing. Antimicrob Agents Chemother 44:3074–3078. Jain SK, Sahu R, Walker LA, Tekwani BL. 2012. A parasite rescue and transformation assay for antileishmanial screening against intracellular Leishmania donovani amastigotes in THP1 human acute monocytic leukemia cell line. J Vis Exp 70:e4054. doi:10.3791/4054. Vanaerschot M, Decuypere S, Downing T, Imamura H, Stark O, De Doncker S, Roy S, Ostyn B, Maes L, Khanal B, Boelaert M, Schönian G, Berriman M, Chappuis F, Dujardin J-C, Sundar S, Rijal S. 2012. Genetic markers for SSG resistance in Leishmania donovani and SSG treatment failure in visceral leishmaniasis patients of the Indian subcontinent. J Infect Dis 206:752–755. Torres DC, Ribeiro-Alves M, Romero GAS, Dávila AMR, Cupolillo E. 2013. Assessment of drug resistance related genes as candidate markers for treatment outcome prediction of cutaneous leishmaniasis in Brazil. Acta Trop 126:132–141. Kaminsky R, Zweygarth E. 1989. Feeder layer-free in vitro assay for screening antitrypanosomal compounds against Trypanosoma brucei brucei and T. b. evansi. Antimicrob Agents Chemother 33:881–885. Yarlett N, Goldberg B, Nathan HC, Garofalo J, Bacchi CJ. 1991. Differential sensitivity of Trypanosoma brucei rhodesiense isolate to in vitro lysis by arsenicals. Exp Parasitol 72:205–215. Raz B, Iten M, Grether-Buhler Y, Kaminsky R, Brun R. 1997. The Alamar Blue® assay to determine drug sensitivity or African trypanosomes (T. b. rhodesiense and T. b. gambiense) in vitro. Acta Trop 68:139–147. Vincent IM, Creek D, Watson DG, Kamleh MA, Woods DJ, Wong PE, Burchmore RJS, Barrett MP. 2010. A molecular mechanism for eflornithine resistance in African trypanosomes. PLoS Pathog 6:e1001204. doi:10.1371/journal.ppat.1001204. Stewart ML, Burchmore RJS, Clucas C, Hertz-Fowler C, Brooks K, Tait A, MacLeod A, Turner CMR, de Koning HP, Wong PE, Barrett MP. 2010. Multiple genetic mechanisms lead to loss of functional TbAT1 expression in drug-resistant trypanosomes. Eukaryot Cell 9:336–343. Maser P, Sutterlin C, Kralli A, Kaminsky R. 1999. A nucleoside transporter from Trypanosoma brucei involved in drug resistance. Science 285:242–244. Carter NS, Barrett MP, de Koning HP. 1999. A drug resistance determinant in Trypanosoma brucei. Trends Microbiol 7:469–471.

151. Susceptibility Test Methods: Parasites n 91. Black CL, Steinauer ML, Mwinzi PNM, Secor WE, Karanja DMS, Colley DG. 2009. Impact of intense, longitudinal retreatment with praziquantel on cure rates of schistosomiasis mansoni in a cohort of occupationally exposed adults in western Kenya. Trop Med Int Health 14:450–457. 92. Guidi A, Andolina C, Ame SM, Albonico M, Cioli D, Haji HJ. 2010. Praziquantel efficacy and long-term appraisal of schistosomiasis control in Pemba Island. Trop Med Int Health 15:614–618. 93. Wang W, Dai J-R, Li H-J, Shen X-H, Liang Y-S. 2010. Is there reduced susceptibility to praziquantel in Schistosoma japonicum? Evidence from China. Parasitology 137:1905– 1912. 94. Cioli D, Botros SS, Wheatcroft-Francklow K, Mbaye A, Southgate V, Tchuenté L-AT, Pica-Mattoccia L, Troiani AR, el-Din SHS, Sabra A-NA, Albin J, Engels D, Doenhoff MJ. 2004. Determination of ED50 values for praziquantel in praziquantel-resistant and -susceptible Schistosoma mansoni isolates. Int J Parasitol 34:979–987. 95. Melman SD, Steinauer ML, Cunningham C, Kubatko LS, Mwangi IN, Wynn NB, Mutuki MW, Karanja DMS, Colley DG, Black CL, Secor WE, Mkoji GM, Loker ES. 2009. Reduced susceptibility to praziquantel among naturally occurring Kenyan isolates of Schistosoma mansoni. PLoS Negl Trop Dis 3:e504. doi:10.1371/journal.pntd. 0000504.

2571

96. Pica-Mattoccia L, Cioli C. 2004. Sex- and stage-related sensitivity of Schistosoma mansoni to in vivo and in vitro praziquantel treatment. Int J Parasitol 34:527–533. 97. Pica-Mattoccia L, Doenhoff MJ, Valle C, Basso A, Troiani A-R, Liberti P, Festuci A, Guidi A, Cioli D. 2009. Genetic analysis of decreased praziquantel sensitivity in a laboratory strain of Schistosoma mansoni. Acta Trop 111:82–85. 98. Liang YS, Coles GC, Doenhoff MJ, Southgate VR. 2001. In vitro responses of praziquantel-resistant and -susceptible Schistosoma mansoni to praziquantel. Int J Parasitol 31:1227–1235. 99. Lamberton PHL, Hogan SC, Kabatereine NB, Fenwick A, Webster JP. 2010. In vitro praziquantel test capable of detecting reduced in vivo efficacy in Schistosoma mansoni human infections. Am J Trop Med Hyg 83:1340–1347. 100. Norton AJ, Gower CM, Lamberton PHL, Webster BL, Lwambo NJS, Blair L, Fenwick A, Webster JP. 2010. Genetic consequences of mass human chemotherapy for Schistosoma mansoni: population structure pre- and postpraziquantel treatment in Tanzania. Am J Trop Med Hyg 83:951–957. 101. Huyse T, Van den Broeck F, Jombart T, Webster BL, Diaw O, Volckaert FAM, Balloux F, Rollinson D, Polman K. 2013. Regular treatments of praziquantel do not impact the genetic make-up of Schistosoma mansoni in northern Senegal. Infect Genet Evol 18:100–105.

Author Index

Volume 1 comprises pages 1 to 1390; volume 2 comprises pages 1391 to 2572.

Abbott, April N., 1379 Abbott, Sharon L., 714 Aberle, Stephan W., 1704 Aguzzi, Adriano, 1859 Ahmed, Abdalla O. A., 2173 Alanio, Alexandre, 2087 Alexander, David C., 441 Anderson, David A., 1584 Araj, George F., 863 Ashbee, H. Ruth, 2965 Atlas, Ronald M., 316, 1955 Atmar, Robert L., 1470

Ciofu, Oana, 773 Clain, Jérôme, 2550, 2563 Coenye, Tom, 838 Conrads, Georg, 967 Conville, Patricia S., 504 Cools, Piet, 813 Copsey, Sarah D., 920 Counihan, Natalie A., 1584 Cox, David L., 1055 Cox, Francis E. G., 2285 Currie, Bart J., 791 Cushion, Melanie T., 2015

Babady, N. Esther, 1498 Bankowski, Matthew J., 1913 Baron, Ellen Jo, 270, 905 Becker, Karsten, 354 Bellini, William J., 1519 Bernard, Kathryn A., 437, 474 Biggs, Beverley-Ann, 2448 Binnicker, Matthew J., 91 Bjarnsholt, Thomas, 773 Bogdan, Christian, 1159 Bopp, Cheryl A., 685, 762 Borman, Andrew M., 2128 Bouyer, Donald H., 1122 Bowen, Michael D., 1660 Brandt, Claudia, 383 Brandt, Mary E., 1935, 1984 Branson, Bernard M., 1436 Brown, Kevin E., 1818 Brown-Elliott, Barbara A., 570, 595 Bruckner, David A., 2357 Bryant, Amy E., 940 Buller, Richard S., 1803 Bush, Karen, 1171

Damon, Inger K., 1828 de Hoog, G. Sybren, 2153, 2173 Deplazes, Peter, 2209 Desmond, Edward P., 1356 Diekema, Daniel J., 106 Doern, Christopher D., 44 Doern, Gary V., 667 Dromer, Françoise, 2087 Dumler, J. Stephen, 873, 1082, 1135 Dunn, James J., 1405

Caliendo, Angela M., 1432 Cama, Vitaliano, 2435 Carpenter, A. Betts, 91 Carroll, Karen C., 29, 940 Carvalho, Maria da Glória Siqueira, 403 Cavling-Arendrup, Maiken, 2255 Chatterjee, Soumya, 2461 Chen, Sharon C.-A., 2030 Chou, Sunwen, 1894 Christensen, Jens Jørgen, 350, 422

Echavarria, Marcela, 1769 Edelstein, Paul H., 887 Elias, Johannes, 635 Escalante, Hermes, 2471 Essig, Andreas, 1106 Facklam, Richard R., 403 Fang, Ferric C., 1379 Farmer, III, J. J., 762 Fields, Patricia I., 685 Finegold, Sydney M., 909 Fitzgerald, Collette, 998 Forman, Michael S., 1599 Forsythe, Stephen J., 714 Franka, Richard, 1633 Frei, Reno, 183 Frosch, Matthias, 635 Fulhorst, Charles F., 1660 Funke, Guido, 474 Gadsby, Naomi J., 1565 Garcia, Lynne S., 2293, 2317

xxix

MCM 11 Edition X : MCM11$CHIX 03-24-15 05:38:01

Page 29

Garcia, Hector H., 2471 Garcia-Hermoso, Dea, 2087 Gärtner, Barbara C., 1738 Gaydos, Charlotte A., 1106 Geißdörfer, Walter, 1159 Gerner-Smidt, Peter, 131 Ghannoum, Mahmoud A., 2188 Ginocchio, Christine C., 1422, 1432, 1783 Gladney, Lori M., 738 Glatzel, Markus, 1859 Gómez, Beatriz L., 2109 Graves, Stephen R., 1150 Gravitt, Patti E., 1783 Guarro, Josep, 2153 Hall, Val, 920 Hanlon, Cathleen A., 1633 Harris, Patricia, 1422 Hayden, Randall T., 1769 Hazen, Kevin C., 1984 Hecht, David W., 1342 Hemarajata, Peera, 238 Heneine, Walid, 1458 Highlander, Sarah K., 226 Hindler, Janet A., 1314 Hodinka, Richard L., 1617, 1718 Hodowanec, Aimee C., 1869 Høiby, Niels, 773 Holfelder, Martin, 44 Horneman, Amy J., 752 Horvat, Rebecca T., 1841 Howell, Susan A., 1984 Huang, Diana G., 1913 Hughes, Laura, 1828 Humphries, Romney M., 1314 Hunsperger, Elizabeth, 1644 Icenogle, Joseph P., 1519 Isham, Nancy C., 2188 Jerome, Keith R., 1687 Jimenez, Juan A., 2471 Johnson, Elizabeth M., 2255 Jones, Jeffrey L., 2373 Jones, Malcolm K., 2479 Jorgensen, James H., 3, 1253

xxx

n

AUTHOR INDEX

Kämpfer, Peter, 813 Kaper, James B., 685 Karlowsky, James A., 1274 Keiser, Jennifer, 2479 Keller, Nancy, 2188 Könönen, Eija, 967 Ksiazek, Thomas G., 1669 Labarca, Jaime, 2357 Lamson, Daryl M., 1536 Landry, Marie Louise, 1432, 1551 LaRocco, Mark T., 1955 Lawson, Andy J., 1013 Le Bras, Jacques, 2550, 2563 Leber, Amy L., 2399 Ledeboer, Nathan A., 667 Leder, Karin, 2529 Lefkowitz, Elliot J., 1393 Leland, Diane S., 1487 Levett, Paul N., 1028 Lewis, II, James S., 1171 Limbago, Brandi M., 1286 Lin, Shou-Yean Grace, 1356 Lindsay, David S., 2425 Lindsley, Mark D., 1955 Lindstrom, Stephen E., 1470 Linscott, Andrea J., 2310 LiPuma, John J., 791 Lockhart, Shawn R., 2223 Loeffelholz, Michael, 217 Lortholary, Olivier, 2087 Lu, Xiaoyan, 1551 Lück, Christian, 887 Lurain, Nell S., 1869 MacCannell, Duncan, 131 Massung, Robert, 1150 Mathis, Alexander, 2209 McAuley, James B., 2373 McGowan, Karin L., 1944 McManus, Donald P., 2479 Mendoza, Leonel, 2196 Meyer, Wieland, 2030 Morrow, Rhoda Ashley, 1687 Moter, Annette, 1159 Munjal, Iona, 120 Nachamkin, Irving, 994, 998 Nagy, Elisabeth, 967 Nataro, James P., 685 Neafie, Ronald C., 2493 Nemec, Alexandr, 813 Nichol, Stuart T., 1669 Noble, Michael A., 169 Nolte, Frederick S., 54 Novak-Weekley, Susan, 2399 Nutman, Thomas B., 2461 O’Donnell, Kerry, 2057 Olson, Victoria A., 1828

MCM 11 Edition X : MCM11$CHIX 03-24-15 05:38:01

Page 30

Orciari, Lillian A., 1633 Ostrowsky, Belinda, 120 Owen, S. Michele, 1436, 1458 Paltridge, Graeme P., 2317 Pang, Xiaoli, 1617 Patel, Robin, 29 Patel, Jean B., 1212 Peacock, Sharon J., 791 Pellett, Philip E., 1754 Perlin, David S., 2236 Petersen, Jeannine M., 738, 851 Petrosino, Joseph F., 226 Petti, Cathy A., 161 Pfaller, Michael A., 3, 106 Pfyffer, Gaby E., 536 Pillay, Allan, 1055 Pitout, Johann, 714 Preiksaitis, Jutta, 1738 Pritt, Bobbi S., 2338 Procop, Gary W., 2493 Puchhammer-Stöckl, Elisabeth, 1704 Radolf, Justin D., 1055 Reller, L. Barth, 15 Reller, Megan E., 1135 Richter, Elvira, 570 Richter, Sandra S., 1212, 1274 Riffelmann, Marion, 838 Robinson, Christine, 1769 Rollin, Pierre E., 1669 Romero, José R., 1536 Rota, Paul A., 131 Ruoff, Kathryn L., 350, 422 Ryan, Norbert, 2448 Schriefer, Martin E., 738, 851, 1037 Schuetz, Audrey N., 1342 Scorpio, Diana G., 873 Secor, W. Evan, 2550, 2563 Seña, Arlene C., 1055 Shafer, Robert W., 1894 Sharp, Susan E., 217, 2310 She, Rosemary C., 161 Sheorey, Harsha, 2448 Shewmaker, Patricia Lynn, 403 Shimizu, Robyn Y., 2293, 2317 Simner, Patricia J., 570 Singh, Kamaljit, 2373 Skov, Robert L., 354 Smith, Jennifer S., 1783 Snyder, James W., 316, 441, 1955 Song, Yuli, 909 Sorrell, Tania C., 2030 Spellerberg, Barbara, 383 Spinler, Jennifer K., 238 Stellrecht, Kathleen A., 1536 Stenger, Steffen, 570 Stevens, Dennis L., 940

Strockbine, Nancy A., 685 Summerbell, Richard C., 2128 Sutton, Deanna A., 2057 Swenson, Jana M., 1286 Switzer, William M., 1458 Tang, Yi-Wei, 1432, 1498 Tarr, Cheryl L., 762 Taylor, Ryan, 1841 Taylor-Robinson, David, 1088 Teixeira, Lúcia Martins, 403 Telford, Sam R., III, 2505 Templeton, Kate E., 1565 Theel, Elitza S., 91 Thompson, George R., III, 2109 Thompson, Kenneth D., 1869 Throckmorton, Kurt, 2188 Tipples, Graham, 1754 Trees, Eija, 131 Turenne, Christine Y., 441 Turnidge, John D., 1246, 1253 Valsamakis, Alexandra, 1432, 1599 van de Sande, Wendy W. J., 2173 Van Horn, Gerald, 1422 Vandamme, Peter A. R., 255, 791 Vaneechoutte, Mario, 613, 813 Versalovic, James, 226, 238 Vilela, Raquel, 2196 Visvesvara, Govinda S., 2387 Vogel, Ulrich, 635 von Eiff, Christof, 354 Waites, Ken B., 1088 Walker, David H., 1122 Wallace, Jr., Richard J., 570, 595 Warnock, David W., 1935, 2223 Wauters, Georges, 613, 813 Weber, Rainer, 2209 Weinstein, Melvin P., 15 Weiss, Louis M., 2425 Weller, Peter F., 2529 Wellinghausen, Nele, 462 Wendt, Constanze, 183 Wengenack, Nancy L., 570 Widmer, Andreas F., 183 Wiedbrauk, Danny L., 5 Wilson, Michael L., 15 Wirsing von König, Carl-Heinz, 838 Witebsky, Frank G., 504 Woods, Gail L., 1356 Xiao, Lihua, 2435 Zaki, Sherif, 1669 Zbinden, Reinhard, 652 Zhang, Sean X., 2057

Subject Index

Volume 1 comprises pages 1 to 1390; volume 2 comprises pages 1391 to 2572.

A0 value, 202 A7 agar, 325 A8 agar, 325, 327 AAC-1 β-lactamase, 1300 Abacavir resistance, 1896–1898 Abacavir, for human immunodeficiency virus (HIV), 1869–1870, 1872 Abbott Architect HIV Ag/Ab Combo, 1444 Abbott HCV EIA, 1607–1608 Abbott Plex ID, 1383–1384 Abbott Prism HCV, 1608 Abbott Prism HTLV-I/HTLV-II, 1463 Abbott RealTime CT/NG assay, 639 Abbott RealTime HBV, 1411 Abbott RealTime HCV Genotype II test, 1605–1606 Abbott RealTime HCV, 1411, 1604–1605, 1610 Abbott RealTime HIV-1, 1411, 1442 Abbott RealTime HPV, 1790, 1793 Abbott RealTime PCR, 1110–1111, 1849 ABC transporters, 2241, 2554–2555 ABCD (amphotericin B colloidal dispersion), 2228–2229 Abdominal abscess anaerobic Gram-negative rods, 972 non-spore-forming, anaerobic, Gram-positive rods, 923 Abdominal cramps Cryptosporidium, 2438 herpes B virus, 1697 macrolides, 1183 metronidazole, 1194 nitrofurantoin, 1196 Abdominal fluid specimens, 276 Abdominal pain adenoviruses, 1772 Anisakis, 2493 arenaviruses, 1673 Blastocystis hominis, 2406 Capillaria philippinensis, 2497 Cyclospora cayetanensis, 2428, 2431 Cystoisospora belli, 2428, 2430 Dientamoeba fragilis, 2413 Diphyllobothrium latum, 2472 Enterobius vermicularis, 2454 filoviruses, 1674 Giardia duodenalis, 2411

herpes simplex virus (HSV), 1689 Hymenolepis nana, 2476 influenza virus, 1471 liver trematodes, 2489 Sarcocystis, 2429 Strongyloides stercoralis, 2457 Taenia saginata, 2473 Trichinella, 2495 varicella-zoster virus, 1704 Aberration, 5 ABI SOLiD system, 70 Abiotrophia antimicrobial susceptibilities, 429–430, 1352 antimicrobial susceptibility testing, 1317, 1325–1326 blood culture, 18 clinical significance, 424 description of genus, 423 direct examination, 425 epidemiology and transmission, 423 identification, 426, 428, 429 interpretation of results, 431 isolation procedures, 425 taxonomy, 422–423 Abiotrophia adiacens, 422 Abiotrophia defectiva, 422, 429, 430, 1325 Abiotrophia elegans, 422, 1325 Abiotrophia para-adiacens, 422 Abiotrophia urinae, 431 Abiotrophia viridans, 429, 430 ABLC (amphotericin B lipid complex), 2228–2229 Abortion Campylobacter, 1000 Chlamydia abortus, 1109 Gram-positive anaerobic cocci (GPAC), 910 Leptotrichia, 974 Trypanosoma cruzi, 2363 Abscess Actinomyces, 922 Actinomyces massiliense, 924 Alcaligenes faecalis, 841 Alistipes, 971 Alloscardovia omnicolens, 925 anaerobic Gram-negative rods, 972, 974 Anaerococcus, 911

xxxi

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : open_odd

Page 31

Bacillus licheniformis, 442 Bacteroides, 970 Brevibacillus parabrevis, 443 Burkholderia, 794 Campylobacter, 1000–1001 Citrobacter, 720 Clostridium, 946, 948 Corynebacterium confusum, 487 Corynebacterium durum, 488 Corynebacterium kroppenstedtii, 490 Corynebacterium mucifaciens, 491 Corynebacterium pyruviciproducens, 491 Corynebacterium simulans, 492 Dialister, 974 Edwardsiella, 721 Finegoldia magna, 911 fungi, 1945, 1947–1948 Fusobacterium, 973 Gram-positive anaerobic cocci (GPAC), 910–911 Helcococcus, 425 Histoplasma capsulatum, 2114 hyaline fungi, 2075–2076 Kerstersia, 841 Klebsiella pneumoniae, 718, 723 Microascus, 2075 Mycobacterium haemophilum, 542 Mycobacterium kansasii, 542 Mycobacterium smegmatis, 598 Mycoplasma, 1093 Nocardia abscessus, 516 Nocardia brasiliensis, 516 Nocardia cyriacigeorgica, 516 Nocardia farcinica, 517 Nocardia nova, 517 Nocardia otitidiscaviarum, 517 Nocardia transvalensis, 518 Nocardia veterana, 518 non-spore-forming, anaerobic, Gram-positive rods, 923 Olsenella, 925 Paenibacillus macerans, 443 Parvimonas micra, 911 Peptostreptococcus anaerobius, 911 phaeohyphomycoses, 2161, 2164 Porphyromonas, 971 Prevotella, 972–973

xxxii

n

SUBJECT INDEX

Abscess (continued) Pseudomonas, 776 Roseomonas, 830 Scopulariopsis, 2075 Selenomonas, 974 Serratia, 720 Slackia exigua, 925 specimen collection, transport, and storage guidelines, 272 Sphingobacterium, 825 Staphylococcus, 357, 360 Streptococcus anginosus group, 387 Sutterella, 974 Trueperella bernardiae, 479 Trueperella pyogenes, 479 Tsukamurella paurometabola, 519 Absidia, 1937, 2087, 2091, 2093 Absidia blakesleeana, 2091 Absidia corymbifera, 2088, 2091 Absidia hyalospora, 2091 Absidia ornata, 2088 Absidia ramosa, 2088 Acalculous cholecystitis, see Cholecystitis Acanthamoeba, 887, 1108, 2387–2395 animal inoculation, 2394 antigen detection, 2392 clinical and laboratory diagnosis, 2392–2393 clinical significance, 2390–2391 collection, handling, and storage of specimens, 2391–2392 culture, 2307, 2393–2394 description of agents, 2387–2388 detection, 2327–2330, 2332 enflagellation experiment, 2393 epidemiology, 2389 evaluation, interpretation, and reporting of results, 2395 isolation procedures, 2393–2394 media for culture, 2315–2316 nucleic acid detection, 2392–2393 serology, 2394 stains for detection, 2312, 2316 storage methods, 166 taxonomy, 2387 treatment, 2394–2395, 2542 Acanthamoeba astronyxis, 2387, 2390 Acanthamoeba castellanii, 2387–2388, 2390 Acanthamoeba culbertsoni, 2387, 2390 Acanthamoeba divionensis, 2387, 2390–2391 Acanthamoeba hatchetti, 2387 Acanthamoeba healyi, 2387, 2390 Acanthamoeba lenticulata, 2387 Acanthamoeba monoxenic culture, 2315 Acanthamoeba polyphaga, 2387, 2390–2391, 2394 Acanthamoeba rhysodes, 2387 Acanthocephalans, 2291 Acanthognatha (phylum), 2291 Acanthoparyphium, 2482 Acanthopodida (order), 2287 Acari/Acarina (subclass), 2507, 2511–2513, 2522 Acaricomes, 354, 361 Acariformes (order), 2511 Acarina, 2511–2513 Accidental myiasis, 2516–2517, 2519 AccuProbe test, 642 Blastomyces dermatitidis, 2118–2119 Coccidioides, 2119 Histoplasma capsulatum, 2118 Mycobacterium, 581–582 Staphylococcus aureus, 365 Acedapsone, 1360

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 32

ACE/RV12 ACE detection, 1575 ACE/RV15 One-Step, 1575 Acervuli, 1939–1940 Acetamide agar, 327 Acetamide hydrolysis test, 316 Acetate ascospore agar, 1959 Acetate differential agar, 327 Acetobacteraceae (family), 820, 829–830 Acetyltransferases, 1217, 1228, 1383 Achaetomium, 2062, 2069, 2075, 2161 Achaetomium strumarium, 2062, 2066, 2155, 2160 Achillurbainiidae (family), 2290 Acholeplasma, 1089–1090 Acholeplasma laidlawii, 1089 Acholeplasmataceae (family), 1089 Acholeplasmatales (order), 1089 Achromatic objectives, 8 Achromobacter, 824 antimicrobial susceptibilities, 844–845 clinical significance, 841 collection, transport, and storage of specimens, 842 description of genus, 839 epidemiology and transmission, 840 evaluation, interpretation, and reporting of results, 845 identification, 843 isolation procedures, 842–843 taxonomy, 838 typing systems, 843–844 Achromobacter aegrifaciens, 838, 841 Achromobacter animicus, 838, 841 Achromobacter anxifer, 838, 841 Achromobacter denitrificans, 630–631, 838, 841 Achromobacter dolens, 838, 841 Achromobacter insolitus, 838, 841, 844 Achromobacter insuavis, 838–839, 841 Achromobacter marplatensis, 838, 841 Achromobacter mucicolens, 838, 841 Achromobacter piechaudii, 630–631, 838, 841, 844 Achromobacter pulmonis, 838–839, 841 Achromobacter ruhlandii, 838, 841 Achromobacter sediminum, 838 Achromobacter spanius, 838, 841, 844 Achromobacter spiritinus, 838, 841 Achromobacter xylosoxidans, 630–631, 838, 840–844, 1226 Acidaminococcaceae (family), 909, 969 Acidaminococcus description of, 909 epidemiology, 910 identification, 913, 916 taxonomy, 909 Acidaminococcus fermentans, 909 Acidaminococcus intestini, 909 Acid-fast stain, 321–322 for Mycobacterium, 321–322, 550–551 for parasitology, 2313 modified for parasites in stool specimens, 2319 modified Kinyoun stain, 1957, 2313 modified Ziehl-Neelsen stain, 2313 Acid-fast trichrome stain, 2314, 2316 Acidovorax characteristics of, 802 clinical significance, 795 description of genus, 792 epidemiology and transmission, 793 identification, 802 taxonomy, 792 Acidovorax delafieldii, 630–631, 792, 802

Acidovorax facilis, 630–631, 802 Acidovorax temperans, 630–631, 792, 802 Acidovorax wautersii, 630–631, 792 Acinetobacter, 614, 617–618, 626–627 antibiotic resistance, 1225, 1227, 1232 antimicrobial susceptibilities, 819, 831, 1174–1178, 1180–1181, 1184, 1186, 1193 antimicrobial susceptibility testing, 1255–1256, 1260, 1279–1280 carbapenemases, 1300 characteristics of species, 814 clinical significance, 813–814 colony morphology, 818 commercial sources of chromogenic agar media for, 326 differentiation of Francisella from, 852 epidemiology and transmission, 813 identification, 815–819 antimicrobial susceptibilities, 819 genotyping and epidemiology, 819 species diversity and, 818–819 isolation procedures, 815 taxonomy, 813 Acinetobacter baumannii antibiotic resistance, 1220, 1224, 1226, 1228–1229 antimicrobial susceptibilities, 1175, 1186– 1187, 1193 antimicrobial susceptibility testing, 1279 characteristics of, 814 clinical significance, 813 colony morphology, 818 as ESKAPE pathogen, 714 identification, 816–819 Acinetobacter beijerinckii, 814, 818 Acinetobacter bereziniae, 814, 818 Acinetobacter boissieri, 818 Acinetobacter calcoaceticus, 814, 818–819 Acinetobacter guillouiae, 814, 817–818 Acinetobacter gyllenbergii, 814, 818 Acinetobacter haemolyticus, 814 Acinetobacter johnsonii, 813–814, 817–818 Acinetobacter junii, 813–814, 818 Acinetobacter lwoffii, 813–814 Acinetobacter nectaris, 818 Acinetobacter nosocomialis, 813–814, 818–819 Acinetobacter parvus, 618, 813–814, 818 Acinetobacter pittii, 813–814, 818–819 Acinetobacter radioresistens, 813–814 Acinetobacter schindleri, 814 Acinetobacter soli, 813–814, 818 Acinetobacter ursingii, 813–814, 818–819 Acne, Propionibacterium acnes and, 923–924 Acquired immune deficiency syndrome (AIDS), see AIDS Human immunodeficiency virus (HIV) AcrAB-TolC RND-type efflux pumps, 1218–1219 Acremonium, 1940, 2071, 2076 antifungal susceptibility testing, 2271 key phenotypic features, 2063 microscopy, 1967, 1969 Acremonium atrogriseum, 2071 Acremonium chrysogenum, 1173 Acremonium egyptiacum, 2063, 2071 Acremonium falciforme, 2060, 2173–2174 Acremonium fusidioides, 2063 Acremonium implicatum, 2063 Acremonium kiliense, 2071, 2177 Acremonium persicinum, 2063 Acremonium recifei, 1967, 2063 Acremonium sclerotigenum, 2063, 2071 Acridine orange stain, 322

SUBJECT INDEX Acrodermatitis chronica atrophicans (ACA), 1041, 1043 Acropetal, 1940 Acrophialophora, 2063, 2069, 2073, 2076 Acrophialophora fusispora, 2063, 2076 Actinobacillosis, 654 Actinobacillus antimicrobial susceptibilities, 662 clinical significance, 654 direct examination, 656 epidemiology and transmission, 653 identification, 658–659 isolation procedures, 656 taxonomy and description of, 652 Actinobacillus actinomycetemcomitans, 1328 Actinobacillus equuli, 652–654, 659 Actinobacillus hominis, 652–653, 659 Actinobacillus lignieresii, 652, 654, 659 Actinobacillus suis, 652–654, 659 Actinobacillus ureae, 652–654, 658–659 Actinobacteria (class), 474, 1159 Actinobacteria (phylum) biochemical characteristics of human Eubacterium-like organisms, 930 identification, 926 taxonomy and description of agents, 920– 921 Actinobaculum, 474, 920–921, 924, 926, 928, 930 antimicrobial susceptibilities, 931 identification, 438 taxonomy, 474–475 Actinobaculum massiliae, 927–928 Actinobaculum schaalii, 920, 923–924 Actinobaculum urinale, 920–921, 923–924, 927–928 Actinomadura chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 513–514 description of genus, 506 identification, 438 microscopy, 521 morphologic characteristics, 507 taxonomy, 505 Actinomadura chibensis, 514 Actinomadura cremea, 514 Actinomadura dassonvillei, 508 Actinomadura latina, 510, 512, 514 Actinomadura madurae, 513–514, 521 Actinomadura nitritigenes, 514 Actinomadura pelletieri, 510, 513–514, 521 Actinomadura sputi, 514 Actinomadura vinacea, 514 Actinomucor, 2088 Actinomucor elegans, 2088 Actinomyces, 474, 480, 920–932 antimicrobial susceptibilities, 931, 1172, 1177, 1183–1184, 1188–1190, 1194, 1348 biochemical characteristics, 928 clinical significance, 922–924 description of agents, 920–921 direct examination, 925–926 epidemiology and transmission, 922 evaluation, interpretation, and reporting of results, 931–932 identification, 438, 926–930 isolation procedures, 926 metronidazole resistance, 1352 specimen collection, transport, and handling, 297 taxonomy, 474–475, 920–921 Actinomyces cardiffensis, 923, 928

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 33

Actinomyces dentalis, 924, 927–928 Actinomyces europaeus, 474, 924, 928 Actinomyces funkei, 923–924, 928 Actinomyces georgiae, 924, 928 Actinomyces gerencseriae, 922–923, 926, 928 Actinomyces graevenitzii, 922, 926, 928 Actinomyces hominis, 924, 928 Actinomyces hongkongensis, 924, 927–928 Actinomyces israelii, 297, 922–924, 926, 928, 1182 Actinomyces johnsonii, 929 Actinomyces massiliense, 920, 923–924, 927–928 Actinomyces meyeri, 923–924, 928 Actinomyces naeslundii, 920, 923–924, 927–929 Actinomyces nasicola, 924, 926–928 Actinomyces neuii, 474, 924, 928 Actinomyces neuii subsp. anitratus, 928 Actinomyces neuii subsp. neuii, 928 Actinomyces odontolyticus, 923, 926, 928, 1163 Actinomyces oricola, 924, 926–928 Actinomyces oris, 928–929 Actinomyces radicidentis, 924, 926, 928 Actinomyces radingae, 474, 923, 928 Actinomyces timonensis, 924, 928 Actinomyces turicensis, 474, 923, 928 Actinomyces urogenitalis, 923, 926, 928 Actinomyces viscosus, 494, 927–928 Actinomycetales (order), 1159 Actinomycetes, aerobic, 504–528 antimicrobial susceptibilities, 526–527 antimicrobial susceptibility testing, 1372–1373 clinical significance, 1372 quality control, 1373 reporting of results, 1373 testing method, 1372–1373 chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 511–520 collection, transport, and storage, 520 colonial morphology, 510 description of genera, 506–511 direct examination, 520–521 epidemiology and transmission, 511 evaluation, interpretation, and reporting of results, 528 identification, 522–526 aerial hyphae, 522 cell wall and cell membrane analysis, 523 colonial morphology, 522 genus assignment, 522 microscopic morphology, 522 molecular identification, 523–526 slide cultures, 522 species assignment, 522–523 isolation procedures, 521–522 microscopic morphology, 512 microscopy, 520–521 molecular identification, 523–526 gene sequencing, 524–525 MALDI-TOF MS, 525–526 PCR with amplicon detection, 523 PCR with REA (restriction endonuclease) analysis, 523–524 proteomics, 525–526 pyrosequencing, 525 morphologic characteristics, 507–508 nucleic acid detection, 521 serologic methods, 526 taxonomy, 504–506

n xxxiii

typing systems, 526 Actinomycetoma, 2176 Aculeata, 2518 Acute dermatolymphangioadenitis (ADLA), 2462, 2465 Acute disseminated encephalomyelitis, measles and, 1520 Acute follicular conjunctivitis, adenoviruses and, 1772 Acute hemorrhagic conjunctivitis adenoviruses, 1772 enterovirus, 1540 specimen selection for, 1541 Acute mesenteric lymphadenitis, adenoviruses and, 1772 Acute respiratory disease (ARD), adenoviruses and, 1769, 1772 Acute respiratory distress syndrome (ARDS), Borrelia and, 1041 Acute toxemic schistosomiasis (Katayama fever), 2484 Acyclovir antiviral susceptibility testing, 1916 Epstein-Barr virus, 1739–1740 herpes B virus, 1697 herpes simplex virus (HSV), 1689, 1695, 1919 herpesviruses, 1882–1883, 1885 human herpesvirus 7 (HHV-7), 1761 varicella-zoster virus, 1706, 1712 Acyclovir resistance, 1917 herpes simplex virus (HSV), 1695, 1894–1895 varicella-zoster virus, 1712, 1895 Additives in blood culture media, 17 medium, 347–348 Adefovir, for hepatitis B virus, 1880–1882, 1900 Adefovir resistance, 1851, 1900, 1917, 1921 Adelophialides, 2070 Adenoid-associated viruses, 1619 Adenolymphangitis, 2464 Adenophorea (class), 2289, 2495, 2497 AdenoPlus test, 1413 Adenoviridae (family), 1398, 1400–1401, 1618, 1769 Adenovirus r-gene, 1775 Adenoviruses, 1769–1778 antigen detection, 1773–1774 clinical significance, 1771–1773 collection, transport, and storage of specimens, 1773 cytopathic effect (CPE), 1775–1776 description of the agent, 1769–1770 detection and identification methods, 1433 direct detection, 1773–1775 electron microscopy, 1770, 1773 epidemiology and transmission, 1770–1771 evaluation, interpretation, and reporting of results, 1777–1778 genome types, 1769, 1775 identification, 1776 immunofluorescence detection in R-Mix cells, 1426 isolation procedures, 1775–1776 microscopy, 1773 nucleic acid detection, 1774–1775, 1778 rapid cell culture, 1426 serologic tests, 1777 specimen collection and handling, 1406–1408

xxxiv

n

SUBJECT INDEX

Adenoviruses (continued) taxonomy, 1769–1770 transport medium for, 1409 treatment, 1777 typing systems, 1776–1777 vaccines, 1772 virion morphology, 1769–1770 Adiaspiromycosis, see also Emmonsia clinical significance, 2115 epidemiology and transmission, 2114 evaluation, interpretation, and reporting of results, 2123 nucleic acid detection, 2117 Adlercreutzia, 920–921 Adult respiratory distress syndrome Anaplasma phagocytophilum, 1139 Ehrlichia chaffeensis, 1138 Adult T-cell leukemia/lymphoma (ATLL), 1460–1462 AdvanSure HPV GenoBlot assay, 1791 AdvanSure real-time PCR HPV test, 1790 Advenella clinical significance, 841 collection, transport, and storage of specimens, 842 description of genus, 839 evaluation, interpretation, and reporting of results, 845 identification, 843 taxonomy, 838 Advenella incenata, 617, 632–633, 838, 841 Advia Centaur HBc, 1848 Advia Centaur HBc IgM, 1848 Advia Centaur HBeAg, 1847 Advia Centaur HBsAg, 1847 Advia Centaur HCV, 1607–1608 Advia Centaur total anti-HBs, 1847 Aedes, 2507 Aegyptianella, 1135–1137 Aegyptianella pullorum, 1136 Aerobic actinomycetes, see Actinomycetes, aerobic Aerobic endospore-forming bacteria, see Endospore-forming bacteria, aerobic Aerococcus, 355 antimicrobial susceptibilities, 430 clinical significance, 424 description of genus, 423 epidemiology and transmission, 423–424 identification, 428, 429 taxonomy, 422–423 Aerococcus christensenii, 424, 427, 428 Aerococcus sanguinicola, 423, 424, 427, 428, 430 Aerococcus urinae, 422–423, 424, 427, 428 Aerococcus urinaehominis, 423, 424, 427, 428 Aerococcus viridans, 422–423, 427, 428 Aeromonadaceae, 752 Aeromonadales, 752 Aeromonas, 752–758, 767 antimicrobial susceptibilities, 757, 1180, 1195 antimicrobial susceptibility testing, 1326 β-lactamase, 1326 clinical significance, 754 collection, transport, and storage of specimens, 754 description of genus, 752–753 direct examination, 754–755 epidemiology and transmission, 753–754 evaluation, interpretation, and reporting of results, 757–758 identification, 755–756 isolation procedures, 722, 755

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 34

members of genus, 753 properties of, 763 serologic tests, 756 taxonomy, 752 Aeromonas allosaccharophila, 753 Aeromonas aquariorum, 752, 757 Aeromonas australiensis, 752, 753 Aeromonas bestiarum, 752, 753, 756 Aeromonas bivalvium, 753 Aeromonas cavernicola, 752, 753 Aeromonas caviae, 752, 753, 754, 755, 756, 757, 768, 1326 Aeromonas culicicola, 752 Aeromonas dhakensis, 752, 753, 754, 757 Aeromonas diversa, 752, 753 Aeromonas encheleia, 753 Aeromonas enteropelogenes, 752 Aeromonas eucrenophila, 752, 753, 756 Aeromonas fluvialis, 752, 753 Aeromonas hydrophila, 752, 754, 756, 757 alkaline peptone water, 327 antibiotic resistance, 1226, 1326 antimicrobial susceptibility testing, 1317, 1326 β-lactamase, 1326 Aeromonas hydrophila subsp. dhakensis, 752 Aeromonas hydrophila subsp. hydrophila, 752, 753, 754 Aeromonas hydrophila subsp. ranae, 753 Aeromonas ichthiosmia, 752 Aeromonas jandaei, 753, 756, 1226 Aeromonas media, 752, 753, 756 Aeromonas molluscorum, 753 Aeromonas piscicola, 753 Aeromonas popoffii, 753, 754 Aeromonas rivuli, 752, 753 Aeromonas salmonicida, 752, 753, 756 Aeromonas salmonicida subsp. achromogenes, 753 Aeromonas salmonicida subsp. masoucida, 753 Aeromonas salmonicida subsp. pectinolytica, 753 Aeromonas salmonicida subsp. salmonicida, 752, 753 Aeromonas salmonicida subsp. smithia, 753 Aeromonas sanarellii, 752, 753 Aeromonas schubertii, 753, 755, 756 Aeromonas simiae, 753 Aeromonas sobria, 752, 753, 757, 1326 Aeromonas taiwanensis, 752, 753 Aeromonas tecta, 753 Aeromonas trota, 752, 753, 755, 756 Aeromonas veronii, 752, 753, 756, 757, 1326 Aeromonas veronii bv. sobria, 752, 753, 754, 757 Aeromonas veronii bv. veronii, 753, 755 Affigene CMV Trender, 1726 Affirm VPIII, 2327 Candida, 1998 Trichomonas vaginalis, 2415 Affymetrix arrays, 71, 144 Aflatoxins, 2188–2189, 2191–2192 AFLP, see Amplified fragment length polymorphism (AFLP) analysis African green monkey polyomavirus (species), 1803 African honeybees, 2518 African swine fever, 1644 African tampan, 2515 African tick bite fever, 1124–1126 African trypanosomiasis, 2357; see also Trypanosoma brucei antiparasitic agent resistance mechanisms, 2555

antiparasitic agent susceptibility testing methods, 2564, 2567 clinical features, 2366 diagnosis, 2367 epidemiology, 2366 treatment, 2367–2368, 2530, 2542–2545 Agar dilution susceptibility testing, 1254, 1257–1258 advantages and disadvantages, 1258 antifungal susceptibility testing, 2272 dilution of antimicrobial agents, 1254 for aminoglycoside resistance in enterococci, 1287–1288 for anaerobic bacteria, 1342–1344 incubation conditions, 1343 inoculation procedure, 1343 inoculum preparation, 1343 interpretation and results reporting, 1343–1344 medium, 1342–1343 quality control, 1343 incubation, 1257 inoculation procedures, 1257 interpretation and reporting of results, 1257–1258 preparation, supplementation, and storage of media, 1257 Agar slide culture plate, 1959 Agaricales (order), 1937 Agaricomycetes (class), 1937 Agglutination assay Brucella, 867–868 Francisella, 859 Pythium insidiosum, 2203 Agglutination reactions, 96–97 Aggregatibacter antimicrobial susceptibilities, 662 antimicrobial susceptibility testing, 1328 clinical significance, 654 direct examination, 656 identification, 658–659 isolation procedures, 656 serotyping, 661 taxonomy and description of, 652 Aggregatibacter actinomycetemcomitans, 229, 297, 652, 654, 656, 658–659, 661, 922 Aggregatibacter aphrophilus, 652, 654, 656, 659, 667, 1321, 1328 Aggregatibacter paraphrophilus, 667 Aggregatibacter segnis, 654, 656, 659, 667, 1321 Agilent 2100 Bioanalyzer, 67 Agilent MassCode PCR system, 145 Agilent Technologies microarrays, 144 Agricultural Select Agent Program, 219 Agrobacterium tumefaciens, 1228 ahpC gene, 1357–1358 Aichi virus, 1618 AIDS, 1436; see also Human immunodeficiency virus coexisting conditions/infections Acanthamoeba, 2391 adenoviruses, 1771 Aspergillus, 2033 Candida, 1993 Cryptococcus, 1993 Cryptosporidium, 2437–2439, 2442 Cyclospora cayetanensis, 2429 Cystoisospora belli, 2428, 2431 Epstein-Barr virus, 1740 Giardia duodenalis, 2409 Histoplasma capsulatum, 2114, 2121 human herpesvirus 6 (HHV-6), 1756, 1760

SUBJECT INDEX human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1762– 1763 Leishmania, 2359 Malassezia furfur, 2146 microsporidia, 2213, 2215 molluscum contagiosum virus, 1831 parvovirus B19, 1820 Pneumocystis, 2018–2020 Sporobolomyces, 1994 sporotrichosis, 2164 Talaromyces marneffei, 2046 Toxoplasma gondii, 2375, 2381 Trichosporon, 1994 HIV disease progression to, 1439 Airway obstruction, Epstein-Barr virus and, 1739 Ajellomyces, 1937, 2110 Ajellomyces capsulatus, 2109 Ajellomyces crescens, 2109 Ajellomyces dermatitidis, 1936, 2109 Ajellomyces duboisii, 2109 Ajellomycetaceae (family), 2109, 2196 Alastrim virus, 1830 Albaconazole, 2230 Echinococcus granulosus, 2476 hyaline fungi, 2077 Albendazole, 2529–2532 adverse effects, 2531–2532 Ascaris lumbricoides, 2451, 2455 Capillaria philippinensis, 2497 Enterobius vermicularis, 2454–2455 Fasciolopsis buski, 2490 filarial nematodes, 2465 Giardia duodenalis, 2412 Gnathostoma, 2498 hookworm, 2455–2456 indications for, 2532 Loa loa, 2468 mechanism of action, 2530 microsporidia, 2216 pharmacokinetics, 2530 Sarcocystis, 2431 spectrum of activity, 2530–2531 Strongyloides stercoralis, 2458 Taenia solium, 2476 Toxocara, 2496 Trichinella, 2495 Trichuris trichiura, 2455, 2459 Albicans ID agar, 1952 Alcaligenaceae (family), 838 Alcaligenes, 615 antimicrobial susceptibilities, 845 clinical significance, 841 collection, transport, and storage of specimens, 842 description of genus, 839 evaluation, interpretation, and reporting of results, 845 identification, 843 taxonomy, 838 Alcaligenes aquatilis, 838–839 Alcaligenes faecalis, 614, 628–629, 632–633, 823, 838–839, 841, 843, 845 Alcaligenes faecalis subsp. faecalis, 839 Alcaligenes faecalis subsp. parafaecalis, 839 Alcaligenes faecalis subsp. phenolicus, 839 Alcaligenes odorans, 839 Alcian blue stain, 1956, 1970 Alcohol antiseptic, 184–185 disinfection with alcohols, 194 hand hygiene, 187 surgical scrub, 187

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 35

Alere Clearview Complete HIV 1/2, 1445 Alere Clearview HIV 1/2 Stat Pak, 1445 Alere Determine HIV 1/2 Ag/Ab Combo, 1445 Alere Triage Micro parasite panel, 2441 Alert CMV Early Complete, 1726 Alert Q-CMV Real Time Complete, 1726 Aleurioconidia, 1939–1940, 1969 Aleuriospore, 1940 Alicyclobacillaceae, 441 Aliivibrio, 762 Alishewanella fetalis, 628–629, 822–823 Alistipes characteristics of genus, 970–971 clinical significance, 971 identification, 976–977 taxonomy, 967–969 Alistipes finegoldii, 967, 971, 976, 978 Alistipes indistinctus, 967 Alistipes obesi, 967, 978 Alistipes onderdonkii, 967, 971, 976, 978 Alistipes putredinis, 967, 978 Alistipes senegalensis, 967, 978 Alistipes shahii, 967, 971, 976, 978 Alistipes timonensis, 967, 976 Alkaline peptone salt broth, 327 Alkaline peptone water, 327 Alkaline phosphatase, 316 Allele-specific hybridization, 67, 69 Allergic bronchopulmonary aspergillosis (ABPA), 1969, 2037, 2044 Allergy aminoglycosides, 1182 aspergillosis, 2032, 2037, 2043–2044 bacitracin, 1197 cephalosporins, 1175 chloramphenicol, 1193 Cladosporium, 2162 clavulanic acid, 1177 cockroaches and, 2513 dust mites, 2517 macrolides, 1183 nitrofurantoin, 1196 penicillins, 1173 polymyxins, 1193 quinolones, 1180 red meat, 2516 rifampin, 1195 rifaximin, 1195 Schizophyllum commune, 2075 sulfonamides, 1192 tetracyclines, 1187 tick bites, 2516 vancomycin, 1189 Alliance for the Prudent Use of Antibiotics, 1214 Alloiococcus, 354–355, 429 Alloiococcus otitis antimicrobial susceptibilities, 371 clinical significance, 361 description of family, 356 direct examination, 361 epidemiology and transmission, 357 identification, 367 isolation procedures, 362 taxonomy, 354 Alloprevotella, 968 Alloprevotella rava, 968 Alloprevotella tannerae, 968, 972 Allopurinol, for Trypanosoma cruzi, 2365 Alloscardovia, 920–921 Alloscardovia omnicolens, 925 Allpahuayo virus, 1669, 1672 ALLPATHs, 233

n xxxv

“All-Species Living Tree” project, 260 Allylamine(s), 2223–2224, 2255–2273 Allylamine resistance, 2239, 2245 Alovudine, for adenoviruses, 1777 Alpha diversity, 227 Alphacoronavirus (genus), 1398, 1565 Alphaherpesvirinae (subfamily), 1398, 1687, 1704 α-Naphthol/KOH, 320 Alphapapillomavirus (genus), 1398, 1783 Alphatorquevirus (genus), 1398 Alphavirus (genus), 1399, 1525, 1644, 1646–1648, 1651, 1652, 1655 Alphaviruses, as biothreat agent, 223–224 Alternaria, 2153, 2155, 2159–2160, 2162, 2165, 2167 antifungal susceptibility testing, 2268– 2269, 2271 sick building syndrome, 2192 Alternaria alternata, 2155, 2159–2160, 2162, 2190 Alternaria chlamydospora, 2162 Alternaria infectoria, 2155, 2159–2160, 2162 Alternaria tenuissima, 2162 Alveolar echinococcosis, 2296 Alveolar hydatid disease, 2471, 2476–2477 Alzheimer’s disease, Chlamydia pneumoniae and, 1109 Amantadine antiviral susceptibility testing, 1916 influenza virus, 1471, 1886–1887 rabies virus, 1641 Amantadine resistance, 1903–1904, 1917, 1921 Amapari virus, 1669, 1671 Amass virus, 1830 Ambler classification system for βlactamases, 1223 Amblyomma, 1138, 2507, 2514–2516 Amblyomma americanum, 2515 Amblyomma cajennense, 2515 Amblyomma hebraeum, 2515 Amblyomma maculatum, 2515 Amblyomma variegatum, 2515 Amdovirus (genus), 1818 Amebae, 2399–2408 collection, transport, and storage of specimens, 2399–2400 description of agents, 2399 direct examination, 2400 epidemiology, transmission, and prevention, 2399 evaluation, interpretation, and reporting of results, 2400 key to identification of intestinal amebae, 2321 microscopy, 2400 nonpathogenic, 2407–2408 pathogenic and opportunistic free-living, 2387–2395 animal inoculation, 2394 antigen detection, 2392 clinical and laboratory diagnosis, 2392– 2393 clinical significance, 2389–2391 collection, handling, and storage of specimens, 2391–2392 culture, 2393–2394 description of agents, 2387–2389 direct examination, 2392 enflagellation experiment, 2393 epidemiology, 2389 evaluation, interpretation, and reporting of results, 2395

xxxvi

n

SUBJECT INDEX

Amebae (continued) isolation procedures, 2393–2394 nucleic acid detection, 2392–2393 permanently stained preparations, 2392 serology, 2394 taxonomy, 2387 treatment, 2394–2395 taxonomy, 2399 Amebiasis, 2324, 2332, 2402–2405 commercial kits for immunodetection of serum antibodies, 2296 diagnostic test sensitivity/specificity, 2404 Amebic dysentery, laboratory tests suggested for, 125 Ameboma, 2403 American Board of Forensic Entomology, 2518 American Society for Microbiology (ASM), biothreat agents and, 218–219, 221– 223 American tick bite fever, 1125 American Trudeau Society (ATS) medium, 327 American trypanosomiasis, 2357, 2362–2365; see also Trypanosoma cruzi clinical features, 2362–2364 detection, 2331, 2364–2365 diagnosis, 2364 epidemiology and transmission, 2362 prevention, 2365 treatment, 2365, 2530, 2545 Trypanosoma rangeli, 2360, 2365 xenodiagnosis, 2307, 2365 American Type Culture Collection, 1424 Amies medium, 48 Amies transport medium with charcoal, 327 Amies transport medium without charcoal, 327 Amikacin, 1181–1182, 1198 Acanthamoeba, 2395 antimicrobial susceptibility testing, 1255, 1260 for Mycobacterium infection, 1358–1360, 1369–1370 Amikacin resistance in Mycobacterium tuberculosis complex, 1356, 1360 Aminocyclitols, 1180–1182 Aminoglycoside(s), 1180–1182 adverse effects, 1181–1182, 1360 antimicrobial susceptibility testing, 1255, 1260 mechanism of action, 1181, 1218–1219, 1359 Mycobacterium, 1358–1360 pharmacology, 1181 spectrum of activity, 1181 Aminoglycoside resistance, 1181, 1218–1220 aminoglycoside-modifying enzymes, 782, 1213, 1220 detection by automated antimicrobial susceptibility testing, 1278 in Enterococcus, 1286–1288 molecular, 1383 due to decreased uptake and altered electrical potential, divalent cations, and efflux, 1219–1220 in Enterococcus, 1278, 1286–1288 agar dilution screening method for detecting, 1287–1288 broth microdilution screening method for detecting, 1288 disk diffusion screening method for detecting, 1288

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 36

phenotypic methods for detecting, 1286–1288 Mycobacterium, 1359–1360 Pseudomonas aeruginosa, 782 ribosome modification, 1220 Streptococcus agalactiae, 1320 Aminoglycoside-modifying enzymes (AMEs), 782, 1213, 1220 3-Aminophenylboronic acid, 1301 4-Aminoquinolines, 2536–2537 8-Aminoquinolines, 2539 Amithiozone activity, 1360 adverse effects, 1360 for Mycobacterium infection, 1360 Amniotic fluid specimen collection, transport, and storage guidelines, 276 parasitology, 2300, 2305 PCR for Toxoplasma gondii, 2380 viruses, 1412 Amodiaquine, 2536–2537, 2564 adverse effects, 2537 mechanism of action, 2536 pharmacokinetics, 2537 spectrum of activity, 2537 Amodiaquine resistance, 2552 Amoebaea (class), 2287 Amoebiasis, see Amebiasis Amoebida (order), 2387 Amoebozoa (phylum), 2287, 2387, 2399 Amoxicillin, 1171–1172, 1198 antimicrobial susceptibility testing, 1255, 1259 with clavulanic acid, 1177, 1198 Gardnerella vaginalis, 498 Amoxicillin resistance, in Haemophilus influenzae, 1322 Amoxicillin-clavulanic acid, 1255, 1259 Amoxicillin-clavulanic acid resistance Bacteroides fragilis group, 1346 Burkholderia pseudomallei, 1325 AmpC assay, Etest, 1300 AmpC β-lactamases, 728–730, 1226 Escherichia coli, 695–696, 1299 tests for, 1299–1300, 1383 Amphotericin B, 2228–2229 antifungal susceptibility testing, 2255–2273 Aspergillus, 2044–2045 Candida, 2004–2005 dimorphic fungi, 2121–2122 eumycotic mycetoma fungi, 2181–2182 Fusarium, 2069 hyaline fungi, 2077 Leishmania, 2361–2362 leishmaniasis, 2564, 2566 mucormycosis, 2089, 2097 Naegleria fowleri, 2395 phaeohyphomycosis, 2167 Pythium insidiosum, 2203 scedosporiosis, 2167 spectrum of activity, 2224, 2228–2229 Talaromyces marneffei, 2048 Trichomonas vaginalis, 2551, 2554–2555 yeast species, MICs for, 2005 Amphotericin B colloidal dispersion (ABCD), 2228–2229 Amphotericin B lipid complex (ABLC), 2228–2229 Amphotericin B resistance, 2229, 2239, 2242–2243 Ampicillin, 1171–1173, 1198 anaerobic bacterial susceptibility percentages, 1351

antimicrobial susceptibility testing, 1255, 1259 Gardnerella vaginalis, 498 with sulbactam, 1177–1178, 1198 Ampicillin resistance Bacteroides fragilis group, 1346 Haemophilus influenzae, 1320–1322 Prevotella, 1347 Ampicillin-sulbactam anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1255, 1259 Bacteroides fragilis group susceptibility percentages, 1350 Ampicillin-sulbactam resistance, in Bacteroides fragilis group, 1346 Amplicor, 1110–1111 Amplicor CT/NG assay, 638 Amplicor HIV-1 Monitor version 1.5, 1442–1443 Amplification contamination control, 78–79 inhibitors, 78, 80 Amplified fragment length polymorphism (AFLP) analysis, 261 Aspergillus, 2043 described, 137–138 Fusarium, 2061 nontuberculous mycobacteria (NTM), slowly growing, 585 reproducibility of, 137 Amplified Mycobacterium tuberculosis direct (AMTD) test, 575–576 Amplified ribosomal DNA restriction analysis (ARDRA), 138 AmpliVue, 74 Amur virus, 1660 Amycolata clinical significance, 514 description of genus, 506 identification, 522 morphologic characteristics, 507 taxonomy, 505 Amycolata autotrophica, 512 Amycolatopsis chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 514 description of genus, 506 identification, 438, 522, 527 morphologic characteristics, 507 taxonomy, 505 Amycolatopsis benzoatilytica, 514 Amycolatopsis orientalis, 510, 514 Amycolatopsis palatopharyngis, 514 AN microplate, for anaerobic Gram-negative rods, 977 ANA MIC panel, 1345 Anaerobic bacteria, see also Anaerobic cocci; Anaerobic Gram-negative rods; Anaerobic Gram-positive rods antimicrobial susceptibilities, 1171–1172, 1174–1175, 1177–1178, 1180, 1185, 1188, 1190–1191, 1193– 1194 antimicrobial susceptibility testing, 1342–1352 agar dilution, 1342–1344 antibiograms, 1352 broth microdilution method, 1344–1345 commercial test methods, 1345–1346 indications for testing, 1349 MIC gradient diffusion method, 1345

SUBJECT INDEX reference test methods, 1342–1345 strategies for testing and reporting of susceptibility data, 1352 suggested agents for testing, 1349 β-lactamase tests, 1345–1346, 1352 identification approaches, 905–907 colony morphology, 906 Gram stain, 905–906 MALDI-TOF MS, 37, 906 pyrosequencing, 907 susceptibility testing, 907 toxigenic culture, 906 molecular detection of antibacterial resistance, 1383 resistance patterns in, 1346–1352 Bacteroides fragilis group, 1346–1347, 1350 Bilophila wadsworthia, 1348 Campylobacter gracilis, 1348 Fusobacterium, 1348 Gram-negative rods, 1348 Gram-positive, non-spore-forming bacilli, 1348 Gram-positive, spore-forming bacilli, 1348 Gram-positive cocci, 1348–1349, 1352 Prevotella and Porphyromonas, 1347 Sutterella wadsworthensis, 1348 specimen choice, collection, transport, and handling, 905 Anaerobic cocci Gram-negative anaerobic cocci (GNAC), 909–916 antimicrobial susceptibilities, 916 clinical significance, 911 description of group, 909 epidemiology, 910 evaluation, interpretation and reporting of results, 916 identification, 913, 916 isolation procedures, 912 taxonomy, 909 Gram-positive anaerobic cocci (GPAC), 909–916 antimicrobial susceptibilities, 913, 916 clinical significance, 910–911 collection, transport, and storage of clinical specimens, 911–912 description of group, 909 direct examination, 912 epidemiology, 909–910 evaluation, interpretation and reporting of results, 916 identification, 912–915 isolation procedures, 912 taxonomy, 909–910 Anaerobic colistin-nalidixic acid (CNA) agar, 327–328 Anaerobic culture specimen processing, 287, 289 suitability of specimens for, 281 Anaerobic Gram-negative rods, 967–985 anaerobic, 967–985 antimicrobial susceptibilities, 983–984 β-lactamase tests, 1302–1303 clinical significance, 969–974 collection, transport, and storage of specimens, 975 description of group, 967–969 direct examination, 975–976 epidemiology and transmission, 969 identification, 976–983 isolation procedures, 976 molecular detection, 975–976

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 37

reporting, interpretation, and reporting of results, 984–985 taxonomy, 967–969 unculturable, 983 Anaerobic Gram-positive rods Clostridium, 940–959 non-spore-forming, 920–932 antimicrobial susceptibilities, 931 clinical significance, 922–925 collection, transport, and storage of specimens, 925 direct examination, 925–926 epidemiology and transmission, 922 evaluation, interpretation, and reporting of results, 931–932 identification, 926–930 isolation procedures, 926 serologic tests, 931 taxonomy and description of agents, 920–922 Anaerobiospirillum characteristics of genus, 970–971 clinical significance, 974 identification, 977, 981–982 taxonomy, 969 Anaerobiospirillum succiniciproducens, 969, 974, 982, 994, 997 Anaerobiospirillum thomasii, 969, 974, 982 Anaerococcus, 909, 911 Anaerococcus hydrogenalis, 909–910, 914–915 Anaerococcus lactolyticus, 909–911, 913–915 Anaerococcus murdochii, 909–910, 913–915 Anaerococcus octavius, 909–910, 914–915 Anaerococcus prevotii, 909–915 Anaerococcus senegalensis, 909 Anaerococcus tetradius, 909–910, 912–915 Anaerococcus vaginalis, 909–915 Anaerofustis, 921–922 Anaerofustis stercorihominis, 922 Anaeroglobus, 909, 916 Anaeroglobus geminatus, 909 AnaeroGRO, 975 Anaeroplasma, 1088–1089 Anaeroplasmataceae (family), 1089 Anaeroplasmatales (order), 1089 Anaerosphaera, 909 Anaerosphaera aminiphila, 910 Anaerostipes, 921 Anaerostipes caccae, 922 Anaerostipes hadrus, 922 Anaerotruncus, 921 Anaerotruncus colihominis, 922, 940 Anal swabs, 2324 Analyte-specific reagents, 54, 82 Anamorph, 1936–1938, 1940, 2153 Anaphylaxis bee stings, 2518 chloramphenicol, 1193 penicillins, 1173 polymyxins, 1193 scorpion venom, 2520 tetracyclines, 1187 Anaplasma, 1135–1145 antimicrobial susceptibilities, 1144 clinical significance, 1083, 1138–1139 collection, transport, and storage of specimens, 283, 1139–1140 description of genus, 1135 diagnostic tests, 1084 epidemiology and transmission, 1083, 1136–1138 evaluation, interpretation, and reporting of results, 1144–1145 features of, 1137

n xxxvii

laboratory confirmation, 1142–1144 phylogenetics, 1136 taxonomy, 1135 Anaplasma bovis, 1136–1137 Anaplasma centrale, 1137 Anaplasma marginale, 1135–1137 Anaplasma ovis, 1137 Anaplasma phagocytophilum, 1130 antigen detection, 1142–1143 antimicrobial susceptibilities, 1144 arthropod vector, 2507 biosafety, 1143 description of, 1135 diagnostic tests, 1084 direct examination, 1142–1143 epidemiology and clinical diseases associated with, 1083 evaluation, interpretation, and reporting of results, 1144–1145 features of, 1137 human granulocytic anaplasmosis (HGA), 1138–1139, 1142–1145 identification, 1143 IFA, 1143–1144 isolation procedures, 1143 laboratory confirmation, 1142–1144 microscopy, 1141–1142 nucleic acid detection, 1143 phylogenetics, 1136 serologic tests, 1084, 1143–1144 Anaplasma platys, 1136–1137 Anaplasmataceae (family), 1135–1136 Ancyclostoma, 2329, 2332, 2501 Ancyclostoma caninum, 2531 Ancyclostoma duodenale, 2323, 2454–2456 clinical significance, 2456 description, 2454 eggs, 2454 larvae, 2454 worms, 2454 diagnosis, 2456 epidemiology and prevention, 2454 taxonomy, 2454 transmission and life cycle, 2456 treatment, 2455–2456 Ancyclostomatidae (family), 2289, 2454 Ancyclostomatoidea (superfamily), 2289, 2454 Ancylistaceae (family), 2087, 2102 Andes virus, 1661–1662 Andromas system, 2042 Anelloviridae (family), 1398, 1400–1401 Anemia chloramphenicol, 1193 Epstein-Barr virus and, 1739 hookworm, 2456 iron deficiency, 2456 Leishmania, 2359 linezolid, 1191 nitrofurantoin, 1196 parvovirus B19, 1819–1820 pernicious, 2472–2473 sulfonamides, 1192 Trichuris trichiura, 2459 Trypanosoma brucei, 2366 viruses, specimens and methods for detection of, 1406 Aneurinibacillus, 438 Anexic culture, amebae, 2393–2394 Angioedema Loa loa, 2467 Mansonella, 2468 Angiostrongyliasis, 2498–2499 Angiostrongylidae (family), 2289

xxxviii n

SUBJECT INDEX

Angiostrongylus, 2498–2499 Angiostrongylus cantonensis, 2500, 2531 Anidulafungin, 2228 antifungal susceptibility testing, 2255–2273 Aspergillus, 2044 dimorphic fungi, 2122 eumycotic mycetoma fungi, 2181–2182 spectrum of activity, 2224 Anidulafungin resistance, 2239 Animal inoculation amebae, pathogenic and opportunistic free-living, 2394 Arenaviridae (family), 1678 Echinococcus multilocularis, 2307 Leishmania, 2361 parasitology, 2307 Trypanosoma brucei, 2367 Trypanosoma cruzi, 2365 Anisakiasis, 2495 Anisakidae (family), 2289, 2493 Anisakis, 2493–2495 clinical significance, 2493 description, 2493 direct examination by microscopy, 2493–2494 epidemiology, transmission, and prevention, 2493 serologic tests, 2493, 2495 treatment, 2495 Anisakis simplex, 2495 Anncaliia, 2209–2211, 2328, 2330 Anncaliia algerae, 2209–2210, 2213, 2215 Anncaliia connori, 2209–2210, 2213 Anncaliia vesicularum, 2209–2210, 2213 Annellides, 1940, 2058 Annellidic conidiogenesis, 1940 Annelloconidia, 1940, 1969 Annellospore, 1940 Anocentor, 2514 Anogenital human papillomavirus, 1784–1785 Anopheles, 2338–2339, 2507 Anopleura (order), 2507, 2510, 2522 Anoplocephalidae (family), 2291 Anorexia arenaviruses, 1673–1674 Balantidium coli, 2417 Dientamoeba fragilis, 2413 filoviruses, 1674 Strongyloides stercoralis, 2457 Trichuris trichiura, 2459 Anterior uveitis, Onchocerca volvulus and, 2466 Anthelminthic agents, 2529–2536; see also specific agents benzimidazoles, 2529–2532 diethylcarbamazine, 2534–2535 ivermectin, 2533–2534 nitazoxanide, 2535–2536 praziquantel, 2532–2533 pyrantel pamoate, 2535 Anthrax, 442, 444–445, 1324; see also Bacillus anthracis biothreat agent, 221 collection, transport, and storage of specimens bioterrorism-related, 446 specimens from animals suspected of having anthrax, 447 specimens from patients suspected of having anthrax, 446–447 cutaneous, 125, 444 gastrointestinal, 444

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 38

inhalational, 444–445 injectional, 445 laboratory tests suggested for, 125 transmission and disease, 221 treatment, 455 vaccine, 447 Anthrax Vaccine Adsorbed, 447 Anthrax Vaccine Precipitated, 447 Anthraxin, 455 Antibacterial agents, 1171–1199; see also Antimicrobial agents; specific drugs aminoglycosides and aminocyclitols, 1180– 1182 antimicrobial susceptibility testing dilution of agents, 1258, 1262 interpretive criteria (breakpoints), 1259–1261 selecting for routine susceptibility testing, 1247–1248, 1254–1256, 1349 bacitracin, 1196–1197 β-lactamase inhibitors, 1177–1178 carbapenems, 1172, 1176–1177 cephalosporins, 1173–1176 chloramphenicol, 1193–1194 concentrations in serum, 1197–1199 fosfomycin, 1196 glycopeptides and lipopeptides, 1187–1189 ketolides, 1184–1185 lincosamides, 1185 macrolides, 1182–1183 metronidazole, 1194 monobactams, 1172, 1176 mupirocin, 1197 nitrofurantoin, 1195–1196 oxazolidinones, 1190–1191 penicillins, 1171–1173 polymyxins, 1192–1193 quinolones, 1178–1180 retapamulin, 1197 rifamycins, 1194–1195 streptogramins, 1189–1190 sulfonamides and trimethoprim, 1191–1192 tetracyclines and glycylcyclines, 1185–1187 Antibacterial resistance ability to detect resistance by automated antimicrobial susceptibility testing, 1278–1280 aminoglycoside resistance in Enterococcus, 1278 carbapenem resistance, 1279–1280 ESBL-producing Enterobacteriaceae, 1279 glycopeptide susceptibility reduction in staphylococci, 1279 in Gram-negative bacteria, 1280 in Gram-positive bacteria, 1279 inducible clindamycin resistance, 1279 linezolid resistance in enterococci and staphylococci, 1278 oxacillin resistance in staphylococci, 1278 penicillin resistance in staphylococci, 1278 Streptococcus resistance, 1279 vancomycin resistance in enterococci, 1278 anaerobic bacteria, resistance patterns in, 1346–1352 Bacteroides fragilis group, 1346–1347, 1350 Bilophila wadsworthia, 1348

Campylobacter gracilis, 1348 Fusobacterium, 1348 Gram-negative rods, 1348 Gram-positive, non-spore-forming bacilli, 1348 Gram-positive, spore-forming bacilli, 1348 Gram-positive cocci, 1348–1349, 1352 Prevotella and Porphyromonas, 1347 Sutterella wadsworthensis, 1348 emergence and spread, 1213–1214 Enterobacteriaceae AmpC cephalosporinases, 728–730 β-lactamases, 727–728 carbapenemases, 728, 730 extended-spectrum β-lactamase (ESBL), 722, 727–729 inoculum effects, 1212 molecular detection, 1379–1385 aminoglycoside, 1383 anaerobic bacteria, 1383 β-lactamases in Gram-negative bacteria, 1383–1384 beta-lactam-resistant pneumococci, 1383 ceftriaxone-resistant Neisseria gonorrhoeae, 1383 fluoroquinolones, 1383 linezolid, 1385 methicillin-resistant Staphylococcus aureus, 1380–1382 mupirocin, 1385 mycobacteria, 1385 resistance targets, 1380–1385 technology, 1379–1380 trimethoprim, 1385 vancomycin-resistant enterococci, 1381–1382 Mycobacterium tuberculosis complex, 1356– 1361 nonhuman niches, 1214 nosocomial, 1214 overview, 1212 phenotypic methods for detecting, 1286–1303 direct tests for β-lactamases, 1302–1303 in Enterobacteriaceae, 1287, 1298–1302 in enterococci, 1286–1289 in staphylococci, 1287, 1289–1297 in streptococci, 1297–1298 Pseudomonas aeruginosa, 781–782 relativity of, 1212 tolerance, 1212–1213 Antibacterial resistance mechanisms, 1212–1235 biochemical mechanisms, 1217–1218 efflux pumps, 1218–1219 modification of the antibiotic, 1217– 1218 modification of the target molecule, 1218 restricted access to target, 1218 for antimicrobial classes, 1218–1235 aminoglycoside resistance, 1218–1220 β-lactam resistance, 1220–1228 chloramphenicol resistance, 1228–1229 daptomycin resistance, 1229 glycopeptide resistance, 1229–1230 ketolide resistance, 1231 linezolid resistance, 1230–1231 macrolide resistance, 1231 metronidazole resistance, 1231–1232 mupirocin resistance, 1235 nitrofurantoin resistance, 1232

SUBJECT INDEX polymyxin resistance, 1232 quinolone resistance, 1232–1233 quinupristin-dalfopristin resistance, 1231 rifampin resistance, 1233 tetracycline resistance, 1233–1234 tigecycline resistance, 1234 trimethoprim-sulfamethoxazole resistance, 1234–1235 genetic basis, 1214–1217 acquisition of resistance genes, 1216– 1217 in Mycobacterium tuberculosis complex, 1357–1361 mutation of cellular genes, 1214–1216 Antibiograms, 1352 Antibiotic resistance screens, 1263 Antibiotic-associated diarrhea Clostridium difficile, 944 Clostridium perfringens, 943–944 Antibiotic-resistant bacteria, inactivation of, 196 Antibody detection, fungi, 1969, 1971 Antibody interference, 100 Anticoagulants, 17 Anticomplement immunofluorescence assay cytomegalovirus (CMV), 1728 Epstein-Barr virus, 1743–1745 Anticonvulsant hypersensitivity syndrome, human herpesvirus 6 (HHV-6) and, 1756 Antifolate(s), 2540–2541 Antifolate resistance, 2564 Antifungal agents, 2223–2230; see also specific drugs allyamines, 2223–2224 azoles, 2224–2227 echinocandins, 2227–2228 novel agents in development, 2230 polyenes, 2228–2229 spectrum and extent of activity, 2224 Antifungal resistance, 2236–2247 allyamine resistance, 2239, 2245 azole resistance, 2237–2243 clinical, 2236 echinocandin resistance, 2237, 2239, 2244–2245 flucytosine resistance, 2239, 2245–2246 microbiological, 2236–2237 polyene resistance, 2237, 2239, 2243 primary, 2237 Antifungal susceptibility testing, 2255–2273 agar dilution method, molds, 2272 Aspergillus, 2044–2045 broth macrodilution method molds, 2269, 2270 yeasts, 2262 broth microdilution method dermatophytes, 2271 molds, 2268–2271 yeasts, 2258–2264 clinical breakpoints, 2257–2258 molds, 2270–2271 yeasts, 2263–2264 colorimetric methods molds, 2271–2272 yeasts, 2264–2265 direct testing on blood samples, yeasts, 2267 disk diffusion method molds, 2272 yeasts, 2266 ECV/ECOFF values, 2255, 2257 Etest molds, 2272

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 39

yeasts, 2267 flow cytometry methods, yeasts, 2265– 2266 gradient strip testing molds, 2272 yeasts, 2267 MALDI-TOF MS, yeasts, 2267 molecular methods molds, 2272–2273 yeasts, 2267–2268 Neo-Sensitabs molds, 2272 yeasts, 2267 proteomic methods, yeasts, 2267 quality control, 2262, 2270 rationale, 2255–2256 test principles, 2256–2257 MFC, 2256–2257, 2273 MIC, 2256 Vitek 2 method, yeasts, 2265 wild-type distribution, 2257 Antifungal susceptibility testing media, 1959 Antifungal Susceptibility Testing Subcommittee of the European Committee on Antibiotic Susceptibility Testing (AFSTEUCAST), 1959 Antigen capture ELISA arenaviruses, 1676 dengue virus, 1648 filoviruses, 1676 Antigen capture enzyme immunosorbent assay (EIA) Cryptosporidium, 2439, 2441 human immunodeficiency virus, 1441 Antigen detection, 24 adenoviruses, 1773–1774 amebae, pathogenic and opportunistic free-living, 2392 Anaplasma phagocytophilum, 1142–1143 arenaviruses, 1676 Aspergillus, 1971–1972, 2038–2039 Bartonella, 877 Blastomyces, 1972 Borrelia, 1043 Campylobacter, 1002 Candida, 1972, 1975, 1977, 1996 Chlamydiaceae, 1112–1113 coccidia, 2430 Coccidioides, 1977 coronaviruses, 1570 Cryptococcus, 1977, 1996–1997 Cryptosporidium, 2439 cytomegalovirus (CMV), 1722–1724 dimorphic fungi causing systemic mycoses, 2116 Ehrlichia chaffeensis, 1140 Entamoeba histolytica, 2404, 2406 enteroviruses, 1542 eumycotic mycetoma, 2178 filoviruses, 1676 flagellates, 2408–2409 Francisella, 856 fungi, 1971–1972, 1975, 1977–1978 Fusarium, 2067–2068 gastroenteritis viruses, 1623–1625 Giardia duodenalis, 2411–2412 hantaviruses, 1663 hepatitis A virus, 1591 hepatitis B virus, 1845–1848 hepatitis C virus, 1603 hepatitis E virus, 1591 herpes simplex virus (HSV), 1691–1692 Histoplasma capsulatum, 1977–1978

n xxxix

human bocavirus, 1823 human metapneumovirus, 1510–1511 hyaline fungi, 2076 influenza viruses, 1472–1476 lymphatic filarial nematodes, 2465 melanized fungi, 2164 mumps virus, 1494 Mycoplasma, 1094 parainfluenza virus, 1489 parasitology, 2307–2308, 2327 parvovirus B19, 1821 Plasmodium, 2347–2348 polyomaviruses, 1806–1807 poxviruses, 1833 rabies virus, 1638–1640 respiratory syncytial virus, 1502–1504 rhinoviruses, 1553–1554 Toxoplasma gondii, 2375 transmissible spongiform encephalopathies (TSEs), 1863–1864 transport medium for testing, 1409–1410 Trichomonas vaginalis, 2327, 2415 Tropheryma whipplei, 1162 varicella-zoster virus, 1707 yeasts, 1996–1997 Antigenemia assays for CMV, 1722–1724 Anti-HSV-1 and anti-HSV-2 ELISA IgG kits, 1693 Antimalarials, 2536–2541 antifolates, 2540–2541 artemisinin derivatives, 2539–2540 atovaquone-proguanil, 2541 quinoline derivatives, 2536–2539 Antimicrobial agents, see also specific drugs antibacterial agents, 1171–1199 disinfectants, see Disinfection/disinfectants for Mycobacterium infection, 1356–1361 amikacin, 1358–1360 aminoglycosides, 1358–1360 amithiozone, 1360 bedaquiline, 1358, 1360 capreomycin, 1358, 1360 clofazimine, 1360 cycloserine, 1360 dapsone, 1360 ethambutol, 1358–1359 ethionamide, 1358, 1360 fluoroquinolones, 1358, 1361 isoniazid, 1357–1358 kanamycin, 1358–1360 linezolid, 1361 macrolides, 1361 PA-824, 1361 p-aminosalicylic acid (PAS), 1361 pyrazinamide, 1358–1359 quinolones, 1361 rifabutin, 1359 rifampin, 1357–1359 rifapentine, 1359 streptomycin, 1358–1360 Antimicrobial stewardship program, 110 Antimicrobial susceptibilities, see specific antimicrobial agents; specific organisms Antimicrobial susceptibility testing ability to detect resistance, 1278–1280 aminoglycoside resistance in Enterococcus, 1278 carbapenem resistance, 1279–1280 ESBL-producing Enterobacteriaceae, 1279 glycopeptide susceptibility reduction in staphylococci, 1279 in Gram-negative bacteria, 1280 in Gram-positive bacteria, 1279

xl

n

SUBJECT INDEX

Antimicrobial susceptibility testing (continued) inducible clindamycin resistance, 1279 linezolid resistance in enterococci and staphylococci, 1278 oxacillin resistance in staphylococci, 1278 penicillin resistance in staphylococci, 1278 Streptococcus resistance, 1279 vancomycin resistance in enterococci, 1278 agar dilution method, 1254, 1257–1258 advantages and disadvantages, 1258 anaerobic bacteria, 1342–1344 dilution of antimicrobial agents, 1254 for aminoglycoside resistance detection in enterococci, 1287–1288 incubation, 1257 inoculation procedures, 1257 interpretation and reporting of results, 1257–1258 preparation, supplementation, and storage of media, 1257 anaerobic bacteria, 1342–1352 agar dilution, 1342–1344 antibiograms, 1352 broth microdilution method, 1344–1345 commercial test methods, 1345–1346 indications for testing, 1349 MIC gradient diffusion method, 1345 quality control, 1343–1345 reference test methods, 1342–1345 strategies for testing and reporting of susceptibility data, 1352 suggested agents for testing, 1349 automated systems, 33–35, 1274–1281 ability to detect resistance, 1278–1280 advantages, 1277–1278 BD Phoenix system, 1277 disadvantages, 1278 MicroScan WalkAway system, 1276– 1277 semiautomated instrumentation for disk diffusion method, 1274–1275 Sensititre ARIS 2X, 1277 VITEK systems, 1275–1276 breakpoint establishment, 1248–1249 clinical and bacteriological response rates, 1248 MIC distributions, 1248, 1253 pharmacokinetics and pharmacodynamics, 1248 broth macrodilution methods, 1258, 1261– 1262 advantages and disadvantages, 1262 dilution of antimicrobial agents, 1258 incubation, 1261 inoculation procedures, 1261 interpretation and reporting of results, 1261–1262 preparation, supplementation, and storage of media, 1258, 1261 broth microdilution method, 1262–1263 ability to detect resistance, 1278–1280 advantages and disadvantages, 1263 anaerobic bacteria, 1344–1345 automated, 1275–1280 breakpoint susceptibility tests, 1263 dilution of antimicrobial agents, 1262 fastidious bacteria, 1315, 1317–1318 for aminoglycoside resistance detection in enterococci, 1288 gradient diffusion method, 1263

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 40

incubation, 1262–1263 inoculation procedures, 1262 interpretation and reporting of results, 1263 manual, 1275 potential agents of bioterrorism, 1316 preparation, supplementation, and storage of media, 1262 resistance screens, 1263 semiautomated, 1275 commercial systems, 1274–1281 confirmatory and supplementary test use, 1250 critical review of results, 1280–1281 dilution methods, 1254–1264 disk diffusion method, 1247, 1264–1269 advantages and disadvantages, 1266– 1267 agar medium for, 1265–1266 aminoglycoside resistance detection in enterococci, 1288 antimicrobial agents recommended for routine testing, 1255–1256 breakpoints, 1259–1261 disks, antimicrobial agent, 1265–1266 fastidious bacteria, 1315, 1317–1318 incubation, 1266 inhibition zone diameter distributions, 1249 inoculation procedure, 1266 international methods, 1268–1269 interpretation and reporting of results, 1254, 1266 interpretive categories, 1249, 1253– 1254 interpretive criteria, 1259–1261, 1265 mupirocin resistance detection in staphylococci, 1297 overview, 1264 quality control, 1267 selection of antibacterial agents for routine testing, 1249 semiautomated instrumentation, 1274– 1275 zone-of-inhibition diameter, 1259–1261, 1265, 1267 D-zone test for clindamycin resistance detection in staphylococci, 1267, 1296–1297 in streptococci, 1267, 1297–1298 error sources, 1270 expert systems, 1250, 1280 fastidious bacteria, 1314–1332 from blood culture bottles, 25 future directions and needs in, 1250 general considerations, 1246–1250, 1253– 1254 hospital infection prevention and, 110– 111 “intermediate” category, 1246, 1249–1250, 1259–1261, 1344 international methods, 1267–1270 diffusion methods, 1268–1269 dilution methods, 1268 EUCAST methods, 1269–1270 interpretive categories, 1246, 1248–1250, 1253, 1259–1261 laboratory information system (LIS), 1274–1277, 1281 manual broth microdilution systems, 1275 MIC breakpoints, 1248–1249, 1253, 1259–1261, 1268 molecular detection, 1249–1250 Mycobacterium, 1356–1373

drugs used for testing, 1361 M. avium complex, 1369–1370 M. kansasii, 1370–1371 M. marinum, 1370–1371 M. tuberculosis complex, 1361–1368 nontuberculous mycobacteria, 1368– 1369 rapidly growing mycobacteria, 1371– 1372 slowly growing nontuberculous mycobacteria, 1371 90-60 rule, 1246 objectives, 1246 problems organisms, 1270–1271 quality control, 1263–1264 anaerobic bacteria, 1343–1345 batch and lot QC, 1264 disk diffusion method, 1267 frequency of testing, 1264, 1267 MIC ranges, 1264 reference strains, 1263–1264, 1267 special disk tests, 1267 zone-of-inhibition diameter ranges, 1267 reporting of results, 1250, 1254 agar dilution method, 1257–1258 anaerobic bacteria, 1343–1345, 1352 antibiograms, 1352 broth macrodilution methods, 1261– 1262 broth microdilution method, 1263 cascade reporting, 1250 disk diffusion method, 1254, 1266 resistance mechanisms and, 1270–1271 selecting a system, 1281 selection of antibacterial agents for testing, 1247–1248, 1254–1256 for anaerobic bacteria, 1349 selection of testing method, 1247, 1253 single-well broth dilution method for clindamycin resistance detection in Staphylococcus, 1267, 1296 for clindamycin resistance detection in Streptococcus, 1267, 1298 for mupirocin resistance detection in Staphylococcus, 1297 for taxonomic identification of Mycobacterium, 601 Antiparasitic agents, 2529–2545 anthelminthic agents, 2529–2536 antiprotozoal agents, 2530, 2535–2545 resistance mechanisms, 2550–2556 African trypanosomiasis, 2555 leishmaniasis, 2554–2555 malaria, 2550–2553 schistosomiasis, 2555–2556 trichomoniasis, 2553–2554 susceptibility testing methods, 2563–2568 African trypanosomiasis, 2564, 2567 future directions, 2567–2568 leishmaniasis, 2564, 2566–2567 malaria, 2563–2566 schistosomiasis, 2564, 2567 trichomoniasis, 2564, 2566 Antiprotozoal agents, 2530, 2535–2545; see also specific agents antimalarials, 2536–2541 benznidazole, 2545 diloxanide furoate, 2541 eflornithine, 2544–2545 iodoquinol, 2541–2542 melarsoprol, 2544 miltefosine, 2542–2543 nifurtimox, 2545

SUBJECT INDEX nitazoxanide, 2535–2536 paromomycin, 2543–2544 pentamidine, 2543 pentavalent antimonials, 2542 suramin, 2544 Antiretroviral drug resistance, in human immunodeficiency virus, 1441, 1447, 1450–1451 Antiretroviral therapy, human immunodeficiency virus, 1440 Antisepsis, definition, 189 Antiseptics, 183–189 alcoholic compounds, 184–185 chlorhexidine, 185 iodophors, 185–186 octenidine, 186 overview, 183–184 PCMX, 186 triclosan, 186 uses of, 186–189 decolonization, 188–189 hand hygiene, 186–187 presurgical skin disinfection, 187–188 surgical hand washing/disinfection, 187 Anti-streptolysin O, 396 Antiterrorism and Effective Death Penalty Act of 1996, 219 Antivenin, scorpion, 2520 Antiviral agent(s), 1869–1887 active against hepatitis B virus, 1880–1882 nucleoside/nucleotide analogues, 1880– 1882 active against hepatitis C virus, 1878–1880 combination therapies, 1880 interferon, 1878–1879 polymerase inhibitors, 1879–1880 protease inhibitors, 1879 ribavirin, 1878–1879 table of agents, 1879 active against herpesviruses, 1882–1886 active against HIV-1 and HIV-2, 1869–1878 entry inhibitors, 1871, 1877 integrase strand transfer inhibitors, 1872, 1877–1878 nonnucleoside reverse transcriptase inhibitors (NnRTIs), 1870, 1873–1874 nucleoside/nucleotide reverse transcriptase inhibitors (NRTIs/ NtRTIs), 1869–1870, 1872–1873 protease inhibitors, 1871, 1874–1875 table of agents, 1870–1872 active against influenza viruses, 1886–1887 M2 protein inhibitors, 1886–1887 neuraminidase inhibitors, 1887 table of agents, 1886 Antiviral resistance causes, 1913–1914 host factors, 1913 patient sociobehavioral influence, 1913–1914 true resistance, 1913 cytomegalovirus (CMV), 1917 hepatitis B virus, 1851–1852, 1899–1900, 1917, 1923–1924 hepatitis C virus, 1900–1903, 1917, 1923– 1924 herpes simplex virus (HSV), 1917 herpesviruses, 1894–1896, 1917–1919

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 41

human immunodeficiency virus (HIV), 1896–1899, 1919–1920, 1923–1924 influenza viruses, 1903–1905, 1917 mechanisms, 1894–1905 varicella-zoster virus, 1917 Antiviral susceptibilities adenoviruses, 1777 arenaviruses, 1681 enteroviruses, 1545 Epstein-Barr virus, 1739, 1740 filoviruses, 1681 hantaviruses, 1665 hepatitis C virus infection, 1609 herpes simplex virus (HSV), 1689, 1695 human herpesvirus 6 (HHV-6), 1760 human herpesvirus 7 (HHV-7), 1762 human herpesvirus 8 (HHV-8), 1764 human metapneumovirus (HMPV), 1512 influenza viruses, 1481–1482 parechoviruses, 1545 polyomaviruses, 1811 rabies virus, 1641 respiratory syncytial virus, 1508 rhinoviruses, 1558 Antiviral susceptibility testing, 1913–1925 clinical indications, 1914 cytomegalovirus (CMV), 1730–1731, 1916, 1918–1919 emerging technologies, 1922, 1924 future directions, 1922, 1924 genotypic assays, 1916–1921 applications, 1918–1921 gene targets, 1917–1918 genotypic platforms, 1916–1918 hepatitis B virus, 1851–1852, 1920–1921 hepatitis C virus, 1921 herpes simplex virus (HSV), 1916, 1919 human immunodeficiency virus (HIV), 1919–1920 influenza, 1916, 1921, 1924 interpretation, 1921–1922 bioinformatics (virtual phenotypes), 1921–1924 genotyping, 1921 phenotypic assays, 1914–1916 dye uptake (DU) test, 1914, 1916 enzyme immunoassay (EIA), 1914, 1916 neuraminidase inhibition assay, 1914, 1916 plaque reduction assay (PRA), 1914– 1916 RVAs, 1915 varicella-zoster virus, 1916, 1919, 1924 Antricola, 2514 Ants, 2518, 2522 Anyplex II MTB/MDR/XDR assay, 583 Anyplex II RV16 detection, 1555, 1575 Anyplex RV16, 1478 Anyplex MTB/NTM assay, 583 Aphanoascus, 2062, 2069 Aphanoascus fulvescens, 2062 Aphthovirus (genus), 1399 API 20 Strep, 428 API 20A, 954, 977 API 20C AUX, 1990, 2001, 2003 API 20E, 34, 453, 614, 745, 768 API 20NE, 614 Bordetella, 843 Burkholderia, 799 Gram-negative nonfermentative rods, 816 Pseudomonas, 778 Stenotrophomonas maltophilia, 801–802 Vibrionaceae, 768

n xli

API 50CH, 487, 494, 927 API 50CHB/E, 453 API CH, 453 API Coryne system, 481, 485, 487–494 API ID 32 GN, 614 API ID 32C, 2001 API NH, 641–642, 658, 676, 678 API Rapid ID 32 Strep system, 392, 409 API Rapid ID 32A, 927 API Rapid NFT, 34 API Staph, 34, 364 API ZYM, 487, 493, 977 Apicomplexa (phylum), 2287, 2338, 2373, 2425, 2435 Apis mellifera adansoni, 2518 APIWEB, 481, 494 Aplastic anemia, see Anemia Apnea, spider envenomation and, 2520 Apochromat objectives, 8 Apoidea, 2518 Aponomma, 2514 Apophysis, 1940 Apophysomyces, 2088, 2091, 2094 Apophysomyces elegans, 1962, 2088, 2094, 2096, 2098 Apophysomyces ossiformis, 2088, 2094 Apophysomyces trapeziformis, 2088, 2094 Apophysomyces variabilis, 2088, 2094 Appendicitis adenoviruses, 1772 Alistipes, 971 anaerobic Gram-negative rods, 972 Ascaris lumbricoides, 2451 Campylobacter, 1001 Enterobius vermicularis, 2454 Fusobacterium, 973 measles, 1521 non-spore-forming, anaerobic, Grampositive rods, 923 Porphyromonas, 971 Sutterella, 974 Appophalus, 2482 Appressorium, 1940 Aptima Combo 2 (AC2), 639 Aptima GC (AG) test, 639 Aptima HCV, 1603 Aptima HCV RNA qualitative test, 1411 Aptima HIV-1 RNA qualitative assay, 75, 1409, 1411, 1441, 1447 Aptima HPV assay, 1414, 1786–1787, 1792, 1793 Aptima HPV genotyping assay, 1794 Aptima transcription-mediated amplification, for Chlamydia trachomatis, 1110–1111 Aptima Trichomonas vaginalis assay, 2327 Aqua-Glo G/C kit, 2441 Aqueous humor, varicella-zoster virus detection in, 1710 Arachnida (class), 1194, 2507, 2511, 2522 Aranaeomorphae, 2520 Araneae (subclass), 2520, 2522 Aravan virus, 1633–1634, 1640 Arboviruses, 1644–1656 antigen detection, 1648 arthropod vector, 2507 biosafety, 1652 characteristics of arboviruses affecting humans, 1645–1646 clinical significance, 1647–1648 collection, transport, and storage of specimens, 1411, 1648 commercial diagnostic tests available, 1649–1651 CPE, 1652

xlii

n

SUBJECT INDEX

Arboviruses (continued) cytopathic effect (CPE), 1652 description of the agent, 1644 detection and identification methods, 1433 direct examination, 1648–1652 enzyme-linked immunosorbent assay (ELISA), 1653–1654 epidemiology and transmission, 1644–1647 evaluation, interpretation, and reporting of results, 1655–1656 identification, 1653 isolation procedures, 1652–1653 microscopy, 1648 nucleic acid detection, 1648–1652 serologic tests, 1653–1655 hemagglutination, 1653 neutralization, 1654–1655 taxonomy, 1644 testing algorithm for human serum specimens from suspected arbovirus infection, 1656 vaccines, 1645–1647 Arcanobacterium, 474 antimicrobial susceptibility testing, 1328 clinical significance, 479 description of genus, 478 epidemiology and transmission, 478 identification, 438, 496 isolation procedures, 480 taxonomy, 474–475 Arcanobacterium bernardiae, see Trueperella bernardiae Arcanobacterium haemolyticum, 478–479, 481, 484, 496 specimen collection, transport, and handling, 300 tetracycline resistance, 1328 vancomycin resistance, 1328 Arcanobacterium pyogenes, see Trueperella pyogenes Archamoebea (class), 2287, 2399 Archiacanthocephala (class), 2291 Architect Anti-HCV, 1607–1608 Architect AUSAB, 1847 Architect Core, 1848 Architect Core-M, 1848 Architect HBsAg, 1847 Architect Syphilis TP, 1071–1072 Arcobacter, 998–1007 antigen detection, 1002 antimicrobial susceptibilities, 1007 clinical significance, 1001 description of agents, 998 identification, 994–995, 997, 1003–1006 isolation procedures, 1003 reservoirs, 999 taxonomy, 998 typing systems, 1006 Arcobacter anaerophilus, 998–999 Arcobacter bivalviorum, 998–999 Arcobacter butzleri, 996, 999, 1001, 1004–1007 Arcobacter cibarius, 999, 1004 Arcobacter cloacae, 998–999 Arcobacter cryaerophilus, 996, 999, 1001, 1004–1005, 1007 Arcobacter defluvii, 998–999 Arcobacter ellisii, 998–999 Arcobacter halophilus, 998–999, 1001 Arcobacter marinus, 998–999 Arcobacter molluscorum, 998–999 Arcobacter mytili, 999, 1001, 1004

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 42

Arcobacter nitrofigilis, 999, 1001, 1004 Arcobacter skirrowii, 996, 998–999, 1001, 1004 Arcobacter suis, 998–999 Arcobacter thereius, 999, 1001, 1004 Arcobacter trophiarum, 998–999 Arcobacter venerupis, 998–999 ARD (acute respiratory disease), adenovirus and, 1769, 1772 ARDRA (amplified ribosomal DNA restriction analysis), 138 ARDS (acute respiratory distress syndrome), Borrelia and, 1041 Arenaviridae (family), 1669–1682 Arenavirus (genus), 1399 Arenaviruses, 1647, 1669–1682 animal inoculation, 1678 antigen detection, 1676 antiviral susceptibilities, 1681 cell culture, 1678 classification, 1399–1400 clinical significance, 1673–1674 collection, transport, and storage of specimens, 1674–1675 postmortem specimens, 1675 safety and security, 1674–1675 shipping, 1675 specimen collection, 1675 description of agents, 1669–1672 detection and identification methods, 1433 direct examination, 1675–1677 electron microscopy, 1675 epidemiology and transmission, 1670 evaluation and interpretation of results, 1681–1682 identification of virus, 1679–1680 immunofluorescence, 1679–1680 isolation procedures, 1677–1678 laboratory tests suggested for, 125 nucleic acid detection, 1677 serologic diagnosis, 1680–1681 ELISA, 1680 IFA test, 1680 neutralization tests, 1680 Western blotting, 1680 taxonomy, 1399–1400, 1669, 1671–1672 typing antisera, 1679 virion morphology, 1401 Argas, 2514 Argasidae (family), 2512, 2514 Argene CINAkit HCMV ppUL83 Rapid Antigenemia, 1724 Argene CMV HHV6,7,8 R-gene, 1726 Argene CMV R-gene, 1726 Argentinian hemorrhagic fever, 1674 Arginine arylamidase, 316 Arginine butyrate, 1740 Arginine dihydrolase, 316–317, 614–615 Arginine hydrolysis, Streptococcus and, 395 ARIS instrument, 34, 1277 Armed spiders, 2520 Armillifer armillatus, 2516 Arrays, see hybridization arrays Arsenic trioxide, HTLVs and, 1461 Artemether, 2540 Fasciola, 2490 Plasmodium, 2349 ARTEMIS Global Antifungal Surveillance Study, 2044, 2238 Artemisinin, 2539–2540, 2564 adverse effects, 2540 mechanism of action, 2540 pharmacokinetics, 2540

Plasmodium, 2349 schistosomes, 2486 spectrum of activity, 2540 Artemisinin resistance, 2551–2552, 2565 Artemisinin-based combination therapies, 2550–2551 Artemisinin-mefloquine, for schistosomes, 2486 Arteritis Lagenidium, 2198, 2204 Pythium insidiosum, 2201 Artesunate, 2540 Fasciola, 2490 herpesviruses, 1886 human herpesvirus 6 (HHV-6), 1756 Plasmodium, 2349 Arthralgia arboviruses, 1647 arenaviruses, 1673 human herpesvirus 8 (HHV-8), 1762 Mansonella, 2468 nitrofurantoin, 1196 rifampin, 1195 rubella, 1526 streptogramins, 1190 Arthritis anaerobic Gram-negative rods, 972 Bartonella, 876 Blastomyces dermatitidis, 2114 Borrelia, 1037, 1041, 1048 Brucella, 865 Burkholderia, 794 Campylobacter, 1000 Chlamydia trachomatis, 1108 Coccidioides, 2114 Dolosigranulum pigrum, 424 Dracunculus medinensis, 2496 Histoplasma capsulatum, 2114 Moraxella, 814 Mycoplasma, 1091, 1093 Pantoea, 719 Porphyromonas, 971 Prevotella, 973 rubella, 1526 Scedosporium boydii, 2177 Sneathia, 974 Staphylococcus, 360 Tannerella forsythia, 972 Tropheryma whipplei, 1160 Yersinia enterocolitica, 742 Arthrobacter, 354, 356 clinical significance, 479 description of genus, 475 epidemiology and transmission, 479 identification, 484, 494 taxonomy, 474–475 Arthrobacter albus, 494 Arthrobacter cumminsii, 479, 494 Arthrobacter oxydans, 494 Arthroconidia, 1939–1940, 1967, 2135, 2138, 2148, 2154 Arthroderma, 1937, 2128 Arthroderma benhamiae, 2131–2132 Arthroderma cajetani, 2129 Arthroderma fulvum, 2130 Arthroderma gertleri, 2133 Arthroderma grubyi, 2128, 2130 Arthroderma gypseum, 2130 Arthroderma incurvatum, 2130 Arthroderma lenticulare, 2133 Arthroderma mirabile, 2129 Arthroderma obtusum, 2130 Arthroderma olidum, 2133 Arthroderma otae, 2129

SUBJECT INDEX Arthroderma persicolor, 2130 Arthroderma quadrifidum, 2133 Arthroderma racemosum, 2130 Arthroderma simii, 2133 Arthroderma uncinatum, 2130 Arthroderma vanbreuseghemii, 2131–2132, 2144 Arthrodermataceae (family), 2109, 2128 Arthrographis, 2063, 2073, 2076 Arthrographis cuboidea, 2073 Arthrographis kalrae, 2063, 2073, 2076 Arthropathy human T-cell lymphotropic viruses (HTLVs), 1461 parvovirus B19, 1819 Arthropods, 2505–2523 as scalars, 2513–2515 cockroaches, 2513, 2515 muscoid flies, 2513 defense against predation by, 2505 delusion or illusion of parasitosis, 2521 direct injury due to, 2515–2521 endoparasitic, 2516–2518 identification of submitted specimens, 2521–2523 key to, 2522 phylogeny, 2505 stinging and biting, 2518–2521 centipedes and millipedes, 2520 Hymenoptera, 2518 scorpions, 2519–2520 spiders, 2520–2521 urticating caterpillars, 2518–2519 vectors, 2505–2513 Acarina, 2511–2513 Diptera, 2505–2508 Hemiptera, 2508–2509 Phthiraptera, 2510–2511 Siphonaptera, 2509–2510 Arthrospore, 1940 Arthus reaction, with atypical measles syndrome, 1520 Articular pain, Mansonella and, 2468 Artus CMV PCR, 1726 Artus HBV PCR, 1849 Artus Influenza A/B RG kit, 1477 Artus M. tuberculosis PCR kit, 575 Artyfechinostomum, 2482 Asaia, 614, 814, 827, 829 Asaia bogorensis, 829 Asaia lannensis, 829 Asaia siamensis, 829 Ascariasis, 2296, 2451 Ascarida (order), 2289, 2448 Ascarididae (family), 2289, 2448, 2495 Ascaridoidea (superfamily), 2289, 2448, 2493 Ascaris lumbricoides clinical significance, 2451 description, 2448–2451 eggs, 2448–2451 larvae, 2451 worms, 2451 detection, 2320, 2323, 2329, 2331 diagnosis, 2451 Dientamoeba transmission and, 2412 epidemiology and prevention, 2451 sputum specimen, 2305 taxonomy, 2448 transmission and life cycle, 2451, 2454 treatment, 2451, 2453, 2455, 2531–2532, 2534–2535 Ascaris suum, 2448 Ascending paralysis

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 43

herpes simplex virus (HSV), 1689 Mycoplasma, 1091 Ascites fluid specimens, 276 Ascocarps, 1938, 1940 Ascoli test, 455 Ascoma, 1940 Ascomycetes homothallic, 2062, 2069, 2075 yeasts ascospores, 2000 description of agents, 1986–1988 taxonomy, 1984–1985 urease-negative, 2001 Ascomycota (phylum), 1936–1938, 1940, 2030, 2057, 2128, 2173–2174, 2196 Ascospore stain, 1956–1957 Ascospores, 1938, 1940, 2000, 2070, 2180 Ascus, 1938, 1940 Asellariales (order), 2087 Aseptate, 1935, 1940 Aseptic meningitis, see also Meningitis enterovirus, 1539–1541 human herpesvirus 6 (HHV-6), 1755 parechovirus, 1539, 1541 specimen selection, 1541 varicella-zoster virus, 1709 Asfaviridae (family), 1644 Ashdown medium, 328 ASHRAE 110 tracer gas test, 173 Aspergilloma, 2037, 2043 Aspergillosis, 1969, 2033, 2036–2037 allergic, 2032, 2037, 2043–2044 antimicrobial susceptibilities, 2044–2045 invasive, 1971, 2031–2033, 2036–2040, 2043–2045 specimens for, 1947 Aspergillus, 1937, 1938, 1939, 1940, 2030–2045, 2164, 2192 aflatoxins, 2188 antibody detection, 1971 antifungal resistance, 2225, 2227, 2237–2238, 2243, 2245–2246, 2255 antifungal susceptibility, 2044–2045, 2224 antifungal susceptibility testing, 2044–2045, 2268–2273 antigen detection, 1971–1972, 2038–2039 (1,3)-β-D-glucan detection, 2038–2039 clinical significance, 1948–1949, 2033, 2036–2037 colony morphology/color, 2031, 2034–2036, 2040–2041 cycloheximide inhibition, 1951, 1955 cyclopiazonic acid, 2189 description, 2031, 2033–2036 direct examination, 2037–2040 epidemiology and transmission, 2031–2033 ergot alkaloids, 2190 evaluation, interpretation, and reporting of results, 2045 identification, 2033, 2040–2043 culture-based, 2040–2042 key, 2033 mass spectrometry, 2042–2043 molecular, 2042 poorly sporulating species or variants, 2042 isolation, 2040 media, 1961 microscopy, 1967, 2033–2038, 2040–2042 nucleic acid detection, 1979, 2039–2040

n xliii

ochratoxins, 2189 serologic tests, 2043–2044 sick building syndrome, 2192 specimen collection, transport, and processing, 1948–1950, 1949, 1950, 2037 staining, 1957 taxonomy, 2030–2033 typing systems, 2043 Aspergillus acidus, 2031 Aspergillus aculeatus, 2031 Aspergillus alabamensis, 2030–2032 Aspergillus alliaceus, 2031–2032 Aspergillus avenaceus, 2031 Aspergillus brasiliensis, 2031–2032 Aspergillus caesiellus, 2031–2032 Aspergillus calidoustus, 2031–2032, 2271 Aspergillus candidus, 2031–2033, 2036 Aspergillus carnus, 2031–2032 Aspergillus chevalieri, 2031–2032 Aspergillus clavatonanicus, 2031–2032 Aspergillus clavatus, 2031–2033 Aspergillus conicus, 2031–2032 Aspergillus deflectus, 2031–2032, 2035 Aspergillus fischeri, 2033 Aspergillus fischerianus, 2031–2032 Aspergillus flavipes, 2031–2034 Aspergillus flavus, 2030–2034, 2037, 2041–2043, 2192 aflatoxins, 2188 antifungal susceptibility testing, 2261, 2271 cyclopiazonic acid, 2189 Aspergillus fumigatiaffinis, 2031–2032 Aspergillus fumigatus, 1997, 2030–2034, 2037, 2039, 2041–2045 antifungal resistance, 2225, 2227, 2237–2239, 2241, 2243 antifungal susceptibility testing, 2255, 2257, 2261, 2268–2273 disinfection, 194 ergot alkaloids, 2190 microscopy, 1973, 1974 nucleic acid detection, 1979 Aspergillus fumisynnematus, 2031–2032 Aspergillus glaucus, 2031–2033, 2036 Aspergillus granulosus, 2031–2032 Aspergillus hollandicus, 2031–2032 Aspergillus janus, 2031 Aspergillus japonicus, 2031 Aspergillus lentulus, 2030–2032, 2034, 2041–2042, 2045, 2237, 2243, 2257 Aspergillus neoniveus, 2189 Aspergillus nidulans, 1938, 1967, 2031–2033, 2035, 2037, 2041, 2043, 2045, 2188, 2228 Aspergillus niger, 2030–2033, 2035, 2037, 2043, 2190, 2271 Aspergillus niveus, 2031–2032 Aspergillus nominus, 2031–2032 Aspergillus ochraceopetaliformis, 2031 Aspergillus ochraceus, 2031–2033 Aspergillus oryzae, 2031–2032, 2189 Aspergillus parasiticus, 2031–2032, 2188 Aspergillus penicillioides, 2031–2032 Aspergillus persii, 2031–2032 Aspergillus reptans, 2031–2032 Aspergillus restrictus, 2031–2033, 2036 Aspergillus rubrobrunneus, 2031–2032 Aspergillus rugulovalvus, 2031–2032 Aspergillus sclerotiorum, 2031–2032 Aspergillus spinosus, 2031–2032 Aspergillus sydowii, 2031–2033 Aspergillus tamari, 2031–2032

xliv

n

SUBJECT INDEX

Aspergillus terreus, 2030, 2032–2033, 2035, 2037, 2043–2045 antifungal resistance, 2243 antifungal susceptibility testing, 2261, 2271 citrinin, 2189 microscopy, 1969 Aspergillus tetrazonus, 2031–2032, 2035 Aspergillus thermomutatus, 2031–2032, 2034 Aspergillus triciti, 2031–2032 Aspergillus tubingensis, 2031–2032 Aspergillus udagawae, 2031–2032, 2034 Aspergillus unguis, 2031–2032 Aspergillus ustus, 2031–2033, 2035, 2041, 2045, 2237, 2243, 2271 Aspergillus vesicolor, 2031–2033, 2036–2037 Aspergillus viridinutans, 2031–2032 Aspergillus wentii, 2031–2033 Asphasmidea (class), 2289 Aspirates Gram stain and plating medium recommendations, 286 parasitology, 2297–2300, 2305–2306 for specimen collection, 270–271 Assassin bugs, 2508 Association of Official Analytical Chemists, 191 Association of Public Health Laboratories, 217 Association of State and Territorial Public Health Laboratory Directors, 1045– 1046 AST, see Antimicrobial susceptibility testing Asteroleplasma, 1088–1089 Asthma adenoviruses, 1771 Aspergillus, 2044 cockroaches, 2513 human metapneumovirus, 1509 respiratory syncytial virus (RSV), 1500 rhinoviruses, 1553 severe asthma with fungal sensitization (SAFS), 2044 Astroviridae (family), 1399–1401, 1618 Astroviruses antigen detection, 1624–1625 cell culture, 1627 clinical significance, 1620 description of agents, 1619 electron microscopy, 1619, 1623 epidemiology and transmission, 1620–1621 evaluation, interpretation, and reporting of results, 1628–1629 molecular detection assays, 1625–1627 PCR, 1626 serologic tests, 1628 specimen collection and handling, 1406 taxonomy, 1618 typing systems, 1628 ASTY, 2265 Asunaprevir resistance, 1902 Ataxia, in tick paralysis, 2516 Atazanavir, for human immunodeficiency virus (HIV), 1871, 1875 Atazanavir resistance, 1897–1898 ATB system, 364 AtheNA HPV study, 1795 AtheNA Multi-Lyte, 1071–1072 AtheNA Multi-Lyte HSV1&2, 1693–1694 Atheroma, Actinomyces, 923 Atherosclerosis Chlamydia pneumoniae, 1109 cytomegalovirus, 1719 Treponema, 1059

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 44

ATLL (adult T-cell leukemia/lymphoma), 1460–1462 Atopic dermatitis, cockroaches and, 2513 Atopobium antimicrobial susceptibilities, 931 clinical significance, 497, 925 enzyme reactions, 929 identification, 930 taxonomy and description, 920–921 Atopobium fossor, 920 Atopobium minutum, 920, 925, 929 Atopobium parvulum, 909–910, 920, 925, 929 Atopobium rimae, 920, 925, 929 Atopobium vaginae, 925, 927, 929, 931 Atovaquone, for Toxoplasma gondii, 2382 Atovaquone resistance, 2551, 2553, 2564 Atovaquone-proguanil, 2349, 2541, 2564 Atovaquone-proguanil resistance, 2565 ATP synthase, mycobacterial, 1360 Atrax, 2520 Atrophic rhinitis, Klebsiella ozaenae and, 718 Auchmeromyia luteola, 2519 Auramine-rhodamine stain, 322 Aureobacterium, see Microbacterium Aureobasidium, 2153, 2159 Aureobasidium pullulans, 2158–2159, 2161, 2163 Auritidibacter description of genus, 477 epidemiology and transmission, 479 identification, 438 taxonomy, 474–475 Auritidibacter ignavus, 477, 484 Australian bat lyssavirus, 1633–1634 Austrobilharzia, 2480 Autism, 905, 974, 1521 Autoclave, 174, 203–204 Autoinoculation, HPV, 1783 Automated blood culture systems, 20–22 Automation automated specimen processing, 48–49 digital imaging, 51 evaluation and selection criteria for systems, 51–52 costs, 52 productivity, 51–52 quality control, 52 reliability, stability, and durability, 52 safety and hygiene, 52 software applications, 52 technical aspects, 52 future perspectives, 52 historical perspective, 47–48 immunoassays, 91, 102–103 limitations of systems, 51 of molecular assays, 74 molecular automation, 48 organism identification, 48 susceptibility testing, 48 total laboratory automation (TLA), 49–51 Autonomic nerve dysfunction, herpes simplex virus (HSV) and, 1689 Autopsy samples, 305–306 AutoSCAN-4 system, 32, 364 Auxacolor 2, 2001, 2003 Avastrovirus (genus), 1618 Avaxim, 1584, 1590 Aveparvovirus (genus), 1818 Average nucleotide identity (ANI), 258, 260 Averyella dalhousiensis, 716, 726 Avian pneumoviruses, 1508 Avibactam, 1178 Avibacterium gallinarum, 655

Avidity assay, 94–95 cytomegalovirus (CMV), 1730 Epstein-Barr virus, 1744 human bocavirus, 1824 human herpesvirus 6 (HHV-6), 1760 parvovirus B19, 1822 Toxoplasma gondii, 2377, 2379–2380 varicella-zoster virus, 1711 Avioq HIV-1 Microelisa system, 1444 Avipolyomavirus (genus), 1803 AVL buffer, 1411 AxSYM Anti-HCV, 1607–1608 AxSYM AUSAB, 1847 AxSYM Core 2.0, 1848 AxSYM Core-M, 1848 AxSYM HBsAg, 1847 Azithromycin, 1182–1183, 1198 antimicrobial susceptibility testing, 1255, 1260 for Mycobacterium, 1361 Toxoplasma gondii, 2382 Azithromycin resistance, 1231, 1323 Azole(s), 2224–2227; see also specific drugs antifungal susceptibility testing, 2255–2273 drug interactions, 2225 for dimorphic fungi, 2121–2122 for entomophthoromycosis, 2099 Leishmania, 2361 mechanism of action, 2224, 2237–2238 pharmacokinetics, 2224–2225 Azole resistance, 2237–2243 Candida, 2006 environmentally acquired, 2239 epidemiology, 2238 intrinsic, 2237 mechanisms, 2238–2243 biofilms, 2242–2243 chromosomal abnormalities, 2242 drug efflux transporters, 2241 drug target modification, 2238–2240 ERG genes, 2238–2242 increasing target abundance, 2240–2241 loss of heterozygosity, 2242 persister cells, 2243 stress adaptation, 2243 regulation of, 2241–2242 structural modeling of resistance, 2239–2240 virulence and, 2242 Azospirillum, 827, 829 AZT, see Zidovudine Aztreonam, 1176, 1198 antimicrobial susceptibility testing, 1255, 1260 with avibactam, 1178 B cells, Epstein-Barr virus infection of, 1738, 1741 B virus, see Herpes B virus Babesia arthropod vector, 2507 blood specimens, 2306 clinical significance, 2351 collection, transport, and storage of specimens, 2351 description of agent, 2349 detection, 2333–2335 direct examination, 2351–2352 epidemiology and transmission, 2349–2351 evaluation, interpretation, and reporting of results, 2352 isolation, 2352

SUBJECT INDEX life cycle, 2350 microscopy, 2351–2352 nucleic acid detection, 2351–2352 Plasmodium falciparum morphology compared, 2352 serologic tests, 2352 stains for detection, 2312–2313 taxonomy, 2338 treatment, 2530 Babesia bovis, 2349 Babesia canis, 2349 Babesia divergens, 2349–2352 Babesia duncani, 2349–2352 Babesia microti, 2349–2352, 2350, 2521, 2523 Babesia venatorum, 2349–2352 Babesiosis, 2349–2352, 2507, 2521, 2523 Bacillaceae, 441, 453 Bacillales (order), 354, 441 Bacillary angiomatosis, 873–874, 876 Bacillary dysentery, 697–699 Bacillary peliosis hepatitis, 876 Bacillus, 441–456, 895 antimicrobial susceptibilities, 455–456, 1177, 1188 antimicrobial susceptibility testing, 1253, 1317, 1326 β-lactamase, 1324, 1326 blood culture contaminant, 18 clinical significance, 442–445 collection, transport, and storage of specimens, 445–447 description of genus, 441 direct examination, 448–450 disinfection, 192–194 epidemiology and transmission, 442 evaluation, interpretation, and reporting of results, 456 identification, 438, 451–454 isolation procedures, 450–451 serologic tests, 454–455 taxonomy, 441 typing, 454 Bacillus abortus, 222 Bacillus alvei, see Paenibacillus alvei Bacillus amyloliquefaciens, 453–454 Bacillus anthracis, 441 antimicrobial susceptibilities, 455 antimicrobial susceptibility testing, 1316, 1324–1325 biothreat agent, 220, 221 characteristics, 220 clinical significance, 444–445 collection, transport, and storage of specimens, 446–447 bioterrorism-related, 446 specimens from animals suspected of having anthrax, 447 specimens from patients suspected of having anthrax, 446–447 colony morphology, 452 direct examination, 448–450 Gram stain, 448 identification, 452–454 isolation of, 450–451 M’Fadyean test, 323, 448–449 polymyxin B-lysozyme-EDTA-thallous acetate agar for, 342 serologic tests, 455 toxins, 445, 455 transmission and disease, 221 typing, 454 vaccines, 447 Bacillus Calmette-Guérin (BCG), 538–539, 555

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 45

Bacillus canis, 222 Bacillus cereus, 441, 443–444, 448–456 antibiotic resistance, 1223, 1226 antimicrobial susceptibilities, 1190 antimicrobial susceptibility testing, 1326 mannitol-egg yolk-polymyxin agar for, 339 specimen collection, transport, and handling, 302–303 Bacillus cereus medium, 328 Bacillus cereus selective agar base, 328 Bacillus circulans, 443, 449, 451–453 Bacillus clausii, 442 Bacillus coagulans, 453 Bacillus cytotoxicus, 441, 443, 445, 453–454 Bacillus firmus, 453 Bacillus idriensis, 443 Bacillus infantis, 443 Bacillus laterosporus, 443, 451 Bacillus lentus, 453 Bacillus licheniformis, 441–443, 451, 453, 1196, 1223 Bacillus megaterium, 441–443, 449, 451–453 Bacillus melitensis, 222 Bacillus mycoides, 441, 443, 453 Bacillus oleronius, 443 Bacillus polymyxa, 1192, 1232 Bacillus pseudomycoides, 441, 443, 454 Bacillus pumilus, 441–443, 449, 452–453 Bacillus sphaericus, see Lysinibacillus sphaericus Bacillus stearothermophilus, see Geobacillus stearothermophilus Bacillus subtilis, 441–442, 449, 451–454 antibiotic resistance, 1228 disinfection, 193–196 Bacillus suis, 222 Bacillus thuringiensis, 441–443, 445, 449–455, 2192 Bacillus toyonensis, 441, 443, 454 Bacillus weihenstephanensis, 441, 443 Bacitracin, 1196–1197 Giardia duodenalis, 2412 Streptococcus, 391 Backache, variola virus and, 1830 BacT/ALERT, 1276 BacT/ALERT blood culture system, 21, 22 BacT/ALERT MB, 1948 Bactec 460 radiometric system, 20–21, 1365 Bactec 460TB system, 553 Bactec 9000 series (BD), 21, 22 Bactec 9050 apparatus, 553 Bactec MGIT 960, 553, 1365–1367 Bactec Myco/F lytic bottles, 21, 1948 Bactec Plus/F bottles, 21 Bacteremia Achromobacter, 841 Acinetobacter, 813 Aerococcus, 424 Alcaligenes faecalis, 841 anaerobic Gram-negative rods, 970, 972 Anaerobiospirillum, 974 Arcobacter, 1001 Arthrobacter, 479 Asaia, 829 Bacillus cereus, 443 Bacillus circulans, 443 Bacillus licheniformis, 442 Bacillus pumilus, 443 Bacillus subtilis, 442 Bacteroides, 970 Bartonella, 873, 876–877 Borrelia, 996 Brevibacterium, 479 Brevundimonas vesicularis, 795 Burkholderia, 794

n xlv

Campylobacter, 997, 1000–1001 Cellulomonas, 479 Cellulosimicrobium, 479 Citrobacter, 720 clostridial, 941–942, 948 Comamonas testosteroni, 795 Corynebacterium amycolatum, 479 Corynebacterium jeikeium, 479 Corynebacterium resistens, 479 Corynebacterium tuberculostearicum, 479 Corynebacterium urealyticum, 479 Delftia acidovorans, 795 Dermabacter hominis, 479 detection, see Laboratory detection of bacteremia and fungemia Dysgonomonas, 655 Enterococcus, 406 Eubacterium, 925 Fusobacterium, 973 Globicatella, 425 Gram-positive anaerobic cocci (GPAC), 910–911 Haemophilus haemolyticus, 670 Haemophilus influenzae, 669–670 Hafnia, 721 Helicobacter, 997, 1017–1018 Herbaspirillum, 997 lactobacilli, 924 Leptospira, 997 Leptotrichia, 973–974 Leuconostoc, 424, 431 Lysinibacillus sphaericus, 443 Microbacterium, 479 Mycoplasma, 1092–1093 Neisseria meningitidis, 637 Neisseria mucosa, 645 Neisseria subflava, 645 non-spore-forming, anaerobic, Gram-positive rods, 923 Ochrobactrum, 824 Olsenella, 925 Paenibacillus polymyxa, 443 Pasteurella, 655 Pediococcus, 424, 431 Photobacterium damselae, 765 Plesiomonas shigelloides, 721 Pseudomonas, 776 Ralstonia, 795 Rhodococcus equi, 519 Robinsoniella peoriensis, 925 Rothia, 479 Rothia mucilaginosa, 361 Salmonella, 701 Selenomonas, 974 Sneathia, 974 Solobacterium moorei, 924 Staphylococcus, 357, 360, 1327 Stenotrophomonas maltophilia, 794 Streptococcus bovis group, 387 Streptococcus mutans group, 387 Streptococcus salivarius group, 387 Tsukamurella tyrosinosolvens, 519 Veillonella, 911 Vibrio, 765–766, 997 Vibrio alginolyticus, 765 Vibrio cincinnatiensis, 766 Vibrio fluvialis, 765 Vibrio furnissii, 765 Vibrio metschnikovii, 766 Weissella, 431 Bacteria classification of bacteria, 255–265 identification by nucleic acid sequencing, 75–76

xlvi

n

SUBJECT INDEX

Bacteria (continued) risk-based classification, 171 storage methods, 166 Bacterial antigen testing, with urine samples, 305 Bacterial Isolate Genome Sequence Database (BIGSdb), 143 Bacterial vaginosis, 2415 anaerobic Gram-negative rods, 969, 972 Atopobium vaginae, 497, 925 Corynebacterium lipophiloflavum, 490 Eggerthella-like taxon, 925 etiologies, usual, 290 Gardnerella vaginalis, 479, 497–498, 925 Gram stains of vaginal smears, 925 Lactobacillus lacking in, 906, 924 Leptotrichia, 497, 974 Mobiluncus curtisii, 925 Mycoplasma, 1092 non-spore-forming, anaerobic, Grampositive rods, 923 Nugent scoring system, 926 placental sample handling for, 292 Porphyromonas, 971 Sneathia, 497 specimen collection, transport, and handling, 296 Tannerella forsythia, 972 Ureaplasma, 1092 Bacteriocidal activity, 1213 Bacteriocidal synergism, 1213 Bacteriological media, 325–347 Bacteriostatic activity, 1212–1213 Bacteroidaceae (family), 967 Bacteroidales (order), 967, 983 Bacteroides antibiotic resistance, 1231, 1234 antimicrobial susceptibilities, 983–984, 1183, 1185–1186, 1346–1347 β-lactamase, 1302, 1342, 1346 characteristics of genus, 970–971 clinical significance, 970–971 epidemiology and transmission, 969 identification, 977–978, 982 isolation procedures, 976 taxonomy and description of genus, 967–969 Bacteroides bile esculin agar, 328 Bacteroides caccae, 978, 984 Bacteroides capillosus, 967, 970 Bacteroides cellulosilyticus, 978 Bacteroides clarus, 978 Bacteroides coprocola, 978 Bacteroides coprophilus, 978 Bacteroides distasonis, 977 Bacteroides dorei, 970, 978 Bacteroides eggerthii, 978, 1350 Bacteroides faecis, 978 Bacteroides finegoldii, 970, 978 Bacteroides fluxus, 978 Bacteroides fragilis antimicrobial susceptibilities, 983–984, 1172–1178, 1180, 1183–1185, 1187, 1190, 1193–1194, 1214, 1346–1347, 1350 antimicrobial susceptibility testing, 1343–1346 β-lactamase, 1346 bile esculin agar with kanamycin for, 328 clinical significance, 970–971 culture of, 922 identification, 977–979, 982 isolation, 976 oxygen tolerance of, 1344

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 46

reference strains, 1343–1344 taxonomy and description, 967–968 thioglycolate bile broth, 345 Bacteroides fragilis group antibacterial resistance patterns, 1342, 1346–1347, 1350 antimicrobial susceptibility testing, 1343–1346, 1352 Bacteroides intestinalis, 978 Bacteroides massiliensis, 971, 977–978 Bacteroides merdae, 977, 984 Bacteroides nordii, 970–971, 977–978, 982 Bacteroides oleiciplenus, 978 Bacteroides ovatus, 967, 970, 978, 983, 1346, 1350 Bacteroides plebeius, 978 Bacteroides pneumosintes, see Dialister pneumosintes Bacteroides pyogenes, 971 Bacteroides salyersiae, 971, 977–978, 982 Bacteroides splanchnicus, 977 Bacteroides stercoris, 978, 984 Bacteroides tectum, 971 Bacteroides thetaiotaomicron, 967, 970, 978, 983–984, 1343–1344, 1346 Bacteroides uniformis, 978, 983–984, 1346, 1350 Bacteroides ureolyticus, see Campylobacter ureolyticus Bacteroides vulgatus, 971, 977–978, 983–984 antibiotic resistance, 1223 antimicrobial susceptibilities, 1346, 1350 Bacteroides xylanisolvens, 978 Bacteroidetes (phylum), 230, 925, 967–969, 983 Baermann technique, 2322, 2457 Baird-Parker medium, 328 Bairnsdale ulcer, 543 BAL, see Bronchoalveolar lavage (BAL) specimen Balamuthia, 2387–2395 stains for detection, 2312, 2316 treatment, 2542 Balamuthia mandrillaris, 2387–2395 animal inoculation, 2394 antigen detection, 2392 clinical and laboratory diagnosis, 2392–2393 clinical significance, 2391 collection, handling, and storage of specimens, 2391–2392 culture, 2393–2394 description of agents, 2389 detection, 2327 enflagellation experiment, 2393 epidemiology, 2389 evaluation, interpretation, and reporting of results, 2395 isolation procedures, 2393–2394 microscopy, 2388, 2390 nucleic acid detection, 2392–2393 permanently stained preparations, 2392 serology, 2394 taxonomy, 2387 treatment, 2395 Balamuth’s aqueous egg yolk infusion medium, 2315 Balanced salt solutions, 1422 Balansia, 2190 Balantidium coli, 2400, 2416–2417 clinical significance, 2417 description, 2416–2417 detection, 2321 direct examination, 2417

epidemiology, transmission, and prevention, 2417 evaluation, interpretation, and reporting of results, 2417 key features, 2416 microscopy, 2417 taxonomy, 2416 treatment, 2417, 2530, 2542 Balantidium suis, 2416 Ballistoconidium, 1941 Ballistospore, 1941 Balneatrix, 813 Balneatrix alpica, 624–625, 826–827 Banking, organism, 113 Bannwarth’s syndrome, 1041 Barbour-Stoenner-Kelly medium, 328 Barmah Forest virus, 1645 Barnesiella, 967 Barnesiella intestinihominis, 967 Barrier filters, 10–11 Bartels respiratory viral detection kit, 1473 Bartels RSV DFA, 1503 Bartholin gland secretions, 276 Bartonella, 873–882 antimicrobial susceptibilities, 881–882, 1180–1181, 1183–1184 clinical significance, 876 collection, transport, and storage of specimens, 877 description of genus, 873 detection in blood, 20 differentiation of Francisella from, 852 direct examination, 877 antigen detection, 877 microscopy, 877 nucleic acid detection, 877 discovery of novel species, 243, 245 epidemiology and transmission, 873–874 evaluation, interpretation, and reporting of results, 882 identification, 879–880 isolation procedures, 877, 879 blood, 877, 879 tissue, 879 serologic tests, 880 species reservoirs, vectors, and hosts, 875 taxonomy, 873 typing systems, 880 zoonotic, 876–877 Bartonella acomydis, 875 Bartonella bacilliformis, 873–877, 879–881, 2507 Bartonella birtlesii, 874–875, 877 Bartonella bovis, 874–875, 877 Bartonella callosciuri, 875 Bartonella capreoli, 873–875, 880 Bartonella chomelii, 874–875 Bartonella clarridgeiae, 873–877, 879–880 Bartonella coopersplainsensis, 875 Bartonella doshiae, 874–875 Bartonella elastic, 873–876 Bartonella elizabethae, 873–875, 877, 881 Bartonella enact, 876 Bartonella grahamii, 873–877 Bartonella henselae, 283, 873–877, 879–881 Bartonella jaculi, 875 Bartonella japonica, 875 Bartonella koehlerae, 873–877, 879 Bartonella melophagi, 873, 875 Bartonella pachyuromydis, 875 Bartonella peromysci, 874–875, 875 Bartonella queenslandensis, 875 Bartonella quintana, 873–877, 879–882, 2507 Bartonella rattaustraliani, 875

SUBJECT INDEX Bartonella rattimassiliensis, 875 Bartonella rochalimae, 873–875, 880 Bartonella schoenbuchensis, 873–875, 880 Bartonella silvatica, 875 Bartonella talpae, 874–875 Bartonella tamiae, 873–875 Bartonella taylorii, 874–875 Bartonella tribocorum, 873–875 Bartonella vinsonii, 873, 881 Bartonella vinsonii subsp. arupensis, 873–876 Bartonella vinsonii subsp. berkhoffii, 874–877, 880 Bartonella vinsonii subsp. vinsonii, 874–875 Bartonella volans, 875 Bartonella washoensis, 873–877 Bartonellaceae (family), 873 Bartonellosis, 2507 Basidiobolaceae (family), 2087, 2102–2103 Basidiobolales (order), 2087, 2102–2103; see also Entomophthoromycosis Basidiobolomycetes (class), 2087 Basidiobolomycosis, 2087, 2099 Basidiobolus, 1937, 1969, 2087, 2098–2103 Basidiobolus ranarum, 2087, 2098–2099, 2102–2103 Basidiocarps, 1938, 1941 Basidioma, 1941 Basidiomycetes hyaline fungi, 2062–2063, 2069–2071, 2075–2076 yeasts description of agents, 1988–1991 taxonomy, 1985 urease-positive, 2000–2001 Basidiomycota (phylum), 1936–1938, 1941, 2057 Basidiospores, 1937–1938, 1941 Basidium, 1937–1938, 1941 Basipetal, 1941 Batch-Learning Self-Organizing Map, 246 Bavovirus (genus), 1617 Bayes’s theorem, 30 Baylisascaris procyonis, 2329, 2332, 2501 Bayou virus, 1661, 1664 BCG (Bacillus Calmette-Guérin), 538–539, 555 BCL card, 453 BCM O157:H7(+) plating medium, 328 BCYEα, 893, 895–897 BD BBL MycoPrep, 2024 BD Bruker MALDI Biotyper, 34; see also Biotyper system BD EpiCenter system for data management, 33–34, 1277 BD GonoGen II test, 641 BD MAX system, 74, 1383–1384, 2024–2025 BD Onclarity HPV test, 1790 BD Phoenix system, 453 antimicrobial susceptibility testing, 1277 automated microbiology system, 33–34 Enterococcus, 411, 1278 Gram-negative bacteria, 1278–1279 Staphylococcus, 364, 1278–1279 Streptococcus, 1320 Streptococcus pneumoniae, 1319 BD ProbeTec SDA assay, 639 BD universal viral transport medium, 1410 BD Veritor, 1474 BD-Kiestra TLA Concept, 49–50 BDxpert system software, 34, 1277 Bear Canyon virus, 1669, 1672 BEAST (Bayesian Evolutionary Analysis and Sampling Trees), 147

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 47

Beauveria, 2063, 2073, 2076 Beauveria bassiana, 2063, 2076 Bedaquiline, 1358, 1360 Bedbugs, 2508–2509, 2515, 2522 Bee venom, 2518 Beef tapeworm, see Taenia saginata Bees, 2518, 2522 Beetles, 2521, 2522 Bell’s palsy herpes simplex virus (HSV), 1689 Mycoplasma, 1091 varicella-zoster virus, 1709 Benomyl, 1950, 2077 Benzidine test (benzidine hydrochloride), 317 Benzimidazole(s), 2529–2532 adverse effects, 2531–2532 Echinococcus multilocularis, 2477 indications for, 2532 mechanism of action, 2530 pharmacokinetics, 2530 spectrum of activity, 2530–2531 Benznidazole, 2545 adverse effects, 2545 mechanism of action, 2545 pharmacokinetics, 2545 spectrum of activity, 2545 Trypanosoma cruzi, 2365 Benzoquinone, 2521 Benzyl benzoate, 2516–2517 Benzyl-arginine arylamidase activity, 616 Bergeyella, 813 Bergeyella zoohelcum, 624–625, 826–829 Bergey’s Manual of Systematic Bacteriology, 258, 263 Berson, Solomon, 91 Bertia, 2502 Besifloxacin, 1178–1179 Beta diversity, 227 Betacoronavirus, 147 Betacoronavirus (genus), 1398, 1565 (1,3)-β-D-Glucan detection Aspergillus, 2038–2039 Cryptococcus, 1996 dimorphic fungi, 2116 fungal detection, 1978 Fusarium, 2067–2068 Pneumocystis, 2024 β-Galactosidase, 317–318 β-Glucuronidase, 318, 391 Beta-hemolytic streptococci antimicrobial susceptibilities, 396–397 antimicrobial susceptibility testing, 1319–1320 colony morphology, 390 identification by Lancefield antigen immunoassays, 391 identification with phenotypic tests and MALDI-TOF MS, 391–392 phenotypic characteristics of, 384 rectal swab screening for, 303 Streptococcus pyogenes, 389 taxonomy, 383 Betaherpesvirinae (subfamily), 1398, 1718, 1754 β-Lactam(s) antimicrobial susceptibility testing, 1255–1259–1260 carbapenems, 1172, 1176–1177 cephalosporins, 1173–1176 monobactams, 1172, 1176 penicillins, 1171–1173 β-Lactam ring cephalosporins, 1173

n xlvii

penicillins, 1171–1172 β-Lactamase Aeromonas, 1326 Bacillus, 1324, 1326 Bacteroides, 1302, 1342, 1346 β-lactamase inhibitors, 1177–1178 Bilophila wadsworthia, 1346, 1348 Burkholderia pseudomallei, 1325 carbapenemases, 1176, 1178, 1223–1226, 1300–1302, 1383 cephalosporinases, 1223, 1227 cephalosporins and, 1173–1175, 1223, 1227 classification Bush group 1, 1226–1227 Bush group 2b penicillinases, 1224– 1226 Bush group 2be, 1224 Bush group 2br, 1225–1226 Bush group 2f, 1225 Bush group 3, 1226 class A β-lactamases, 1224–1226 class B β-lactamases, 1226 class C β-lactamases, 1226–1227 class D β-lactamases, 1227–1228 schemes, 1223 Clostridium clostridioforme, 1348 CTX-M, 1223–1225, 1383–1384 Enterobacteriaceae, 722, 727–730, 1224–1225, 1227–1228 AmpC, 728–730, 1299–1300 carbapenemases, 1176, 1178, 1223– 1226, 1300–1302 extended-spectrum β-lactamase (ESBL), 722, 727–729, 1299 tests for, 1299–1302 Escherichia coli, 695–696, 1224–1227 extended-spectrum β-lactamase (ESBL), 1172, 1173–1175, 1178, 1224–1225, 1254, 1270–1271, 1279 β-lactamase inhibitors, 1178 cephalosporins and, 1173–1175 commercial sources of chromogenic agar media for, 327 Enterobacteriaceae, 324, 327, 722, 727– 729, 1299 Escherichia coli, 695–696 media for detection, 324 molecular detection, 1383 penicillins and, 1172 tests for, 1299 Francisella tularensis, 1325 Fusobacterium, 1348 genetic environment of, 1224 HACEK group, 1328 Haemophilus influenzae, 1302, 1320–1322 IMP type, 1223–1224, 1226, 1383–1384 in Gram-negative bacteria, 1383–1384 inoculum effects, 1212 integrons, 1224 Klebsiella pneumoniae, 1223–1226, 1228, 1299 KPC, 1225, 1279, 1300, 1383–1384 mechanism of action, 1212–1213, 1223 metallo-β-lactamase, 1176, 1178, 1226, 1300–1301, 1346, 1383 Moraxella catarrhalis, 1330 Mycobacterium tuberculosis complex, 1357 Neisseria meningitidis, 1324 OXA (oxacillin hydrolyzing), 1223–1224, 1227–1228, 1300, 1302, 1383–1384

xlviii

n

SUBJECT INDEX

β-Lactamase (continued) Pasteurella, 1331 penicillins and, 1172–1173 Porphyromonas, 1346–1347 Prevotella, 1342, 1346–1347 processing, 1223–1224 Pseudomonas aeruginosa, 781, 1224–1227 resistance, 1220–1228 ROB-1, 1320, 1331 SHV, 1223–1225, 1299, 1383–1384 staphylococcal, 1213–1214, 1289 TEM, 1223–1226, 1299, 1320, 1322 tests for AmpC, 1299–1300 carbapenemases, 1300–1302 direct tests for β-lactamases, 1302–1303 extended-spectrum β-lactamase (ESBL), 1299 in anaerobes, 1345–1346 molecular, 1383–1384 zone edge test, 1289, 1302 VIM type, 1223–1224, 1226, 1383–1384 β-Lactamase inhibitors, 1177–1178 avibactam, 1178 clavulanic acid, 1177 MK7655, 1178 sulbactam, 1177–1178 tazobactam, 1178 β-Lactam resistance, 1220–1228 β-lactamase-mediated resistance, 1223–1228 class A β-lactamases, 1224–1226 class B β-lactamases, 1226 class C β-lactamases, 1226–1227 class D β-lactamases, 1227–1228 common associations of resistance mechanisms, 1215 penicillin-binding protein-mediated, 1212, 1220–1223 acquisition of foreign PBPs, 1221–1222 PBP overexpression, 1221 point mutations, 1222–1223 resistance mutations by recombination with foreign DNA, 1222 pneumococci, 1383 Beta-lysin, 317 Betamethasone, 2515 Betapapillomavirus (genus), 1398, 1783 B-G-Star, 1996 BHI media, for fungi, 1951–1952 BHI-V6 screening agar test method, 1295 Biatriospora mackinnonii, 1968, 2173–2174, 2180–2181 Bibersteinia trehalosi, 655 Bichro-Dubli test, 1986 Bifidobacterium antimicrobial susceptibilities, 931, 1348 clinical significance, 925 identification, 438, 926–927, 930 isolation procedures, 926 taxonomy and description, 920–921 Bifidobacterium adolescentis, 925 Bifidobacterium breve, 925 Bifidobacterium dentium, 925 Bifidobacterium infantis, 920 Bifidobacterium longum, 920, 925 Bifidobacterium scardovii, 925, 927 Bifidobacterium suis, 920 Big Brushy virus, 1669, 1672 BiGGY (bismuth sulfite-glucose-glycine yeast) agar, 1959 Bigyra (phylum), 2406 Bihlvax, 2486 Bile esculin agar, 328

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 48

Bile Bile Bile Bile

esculin agar with kanamycin, 328 esculin azide agar, 328 esculin test, for Streptococcus, 395 fluid specimen collection, transport, and storage guidelines, 276 Bile peritonitis, liver trematodes and, 2489 Bile solubility test, 317, 394–395 Bilharziella, 2480 Biliary tract infection Cyclospora cayetanensis, 2429 Cystoisospora belli, 2428 Bilophila antimicrobial susceptibilities, 1184 characteristics of genus, 970–971 clinical significance, 974 identification, 977 isolation procedures, 976 taxonomy, 969 Bilophila wadsworthia, 969, 974, 976–977, 981 antibacterial resistance patterns, 1348 β-lactamase, 1346, 1348 Binary typing, 143 BinaxNOW malaria test, 2336 BinaxNOW Influenza A&B, 1474 BinaxNOW RSV, 1504 BinaxNOW Staphylococcus aureus test (Alere Scarborough, Inc.), 24 Biocard Adenostick, 1624 Biocard Rotastick, 1624 Biochemical profiles, for organism identification, 29, 30 Biochemical tests, 316–320 Biocides, resistance to, 195–196 BioElisa Syphilis 3.0, 1068 Biofilm Acinetobacter, 817–818 anaerobic Gram-negative rods, 969, 972, 974 azole resistance, 2242–2243 Candida, 2242–2243 echinocandin resistance, 2245 Enterococcus, 406 Fusobacterium, 973 Helicobacter, 1017 Pseudomonas aeruginosa, 774–775, 777, 781 Selenomonas, 974 BioFilmChip microarrays, 72 BioFire FilmArray, see FilmArray Bioinformatics antiviral susceptibility testing, 1921–1924 definitions, 142 BiokitHSV-2 rapid test, 1693–1694 Biolog system, 34, 768 Biological control, with mycoviruses, 2192 bioMérieux Concept FMLA, 50 Bio-Plex 2200 HSV-1 and -2 kit, 1693 Bio-Plex 2200 Syphilis, 1071–1072 Biopsy specimen, see also specific body locations collection and transport, 271, 281 microsporidia, 2215 Mycobacterium, 547–548 parasitology, 2297–2300, 2306 Pneumocystis, 2020, 2021 viruses, 1415–1416 Bio-Rad GS HIV Combo Ag/Ab EIA, 1444 Bio-Rad GS HIV-1/2 PLUS O, 1444 BioRobot EZ1 system, 74 Biosafety Anaplasma phagocyrophilum, 1143 antimicrobial susceptibility testing, 1325 arboviruses, 1652 arenaviruses, 1674–1675 Chlamydiaceae, 1113

dimorphic fungi causing systemic mycoses, 2117 Ehrlichia chaffeensis, 1140 filoviruses, 1674–1675 Francisella tularensis, 855–856 fungi, 1947 Fusarium, 2068 hantaviruses, 1663 influenza viruses, 1479, 1482 prions, 1861–1862 Rickettsia conorii, 1128 specimen collection considerations, 1409 Biosafety cabinets, 172–173, 282–283 Biosecurity, dimorphic fungi causing systemic mycoses and, 2117 BioSign Flu A&B, 1474 Biosynth chromogenic medium for Listeria monocytogenes, 328–329 Bioterrorism, see Biothreat agents Bioterrorism Act, 219 Biothreat agents, 217–224 antimicrobial susceptibility testing, 1316, 1324–1325 botulism toxin, 947 categories and definitions, 219 category A agents, 219, 221–222 anthrax, 221 botulism, 221 hemorrhagic fever viruses, 222 plague, 221 smallpox, 221–222 tularemia, 222 category B agents, 219, 222–224 brucellosis, 222 epidemic typhus, 223 epsilon toxin, 223 food and water safety threats, 224 glanders, 222 melioidosis, 222 psittacosis, 223 Q fever, 222–223 staphylococcal enterotoxins, 223 viral encephalitis, 223–224 characteristics, summary of, 220 Federal Select Agent Program, 219 Laboratory Response Network (LRN), 217–219 mycotoxins, 2192 Rickettsia, 1124 Biotyper system, 34, 48, 72 anaerobic Gram-negative rods, 982 Clostridium, 955 HACEK group, 658, 661 Bipolaris, 1940, 2159–2160, 2268–2269 Bipolaris australiensis, 2160 Bipolaris hawaiiensis, 2160 Bipolaris spicifera, 2160 Bird seed and esculin base medium, 1952 Birdseed agar, 1959, 2000 Birth defects arenaviruses, 1673 rubella, 1526–1527 Bismuth sulfite agar, 329 Bismuth sulfite-glucose-glycine yeast (BiGGY) agar, 1959 Bite wounds, see also Dog bites; Wound infection rabies virus, 1635 specimen collection, transport, and storage guidelines, 272 Biting and stinging arthropods, 2518–2521 centipedes and millipedes, 2520 Hymenoptera, 2518 scorpions, 2519–2520

SUBJECT INDEX spiders, 2520–2521 urticating caterpillars, 2518–2519 Biverticillium, 2045–2046 Bjerkandera adusta, 2077 BK polyomavirus (BK virus), 1803–1812 antigen detection, 1806–1807 antiviral susceptibilities, 1811 cell culture, 1810 clinical significance, 1804–1806 collection, transport, and storage of specimens, 1806 cytopathic effect (CPE), 1810 description of agents, 1803–1804 direct examination, 1806–1810 electron microscopy, 1806 epidemiology and transmission, 1804 evaluation, interpretation, and reporting of results, 1811–1812 in situ hybridization (ISH), 1807–1808 isolation procedures, 1810 nucleic acid amplification tests (NAATs), 1807, 1809–1810 nucleic acid detection, 1807, 1809–1810 serologic tests, 1810 specimen collection and handling, 1407–1408 taxonomy, 1803 BK polyomavirus (species), 1803 BK virus r-gene primers/probe, 1810 BK virus R-gene quantification kit, 1810 BKV primer pair, 1810 Black Creek Canal virus, 1661, 1664 Black grain mycetoma, 1968, 2174, 2176, 2183 Black piedra, 2148 Black yeasts, 2004 Blackflies, 2505–2506, 2515 Blackfly fever, 2506, 2515 Blackwater fever, 2341 Bladder specimens, for parasitology, 2328, 2330 BLAST (Basic Local Alignment Search Tool), 143 Blastic conidiogenesis, 1939–1940, 1941 Blastoconidium, 1941 Blastocystea (class), 2287, 2406 Blastocystida (order), 2406 Blastocystis, 2320, 2406–2407 Blastocystis hominis, 2400, 2406–2407 clinical significance, 2406 description, 2406 diagnosis, 2406–2407 direct examination, 2406–2407 epidemiology, transmission, and prevention, 2406 evaluation, interpretation, and reporting of results, 2407 microscopy, 2406 serologic tests, 2406–2407 taxonomy, 2406 treatment, 2407, 2542–2544 trophozoites and cysts, 2401–2402 Blastomyces, 1937, 1939, 1969, 1972, 2109–2111, 2116 Blastomyces dermatitidis, 895, 1935, 1936, 1938, 1939, 2109–2123 antifungal susceptibilities, 2121–2122, 2224 antigen detection, 2116 biosafety, 2117 clinical significance, 2114 culture for mold phase, 2117 culture for yeast phase, 2118 description of agents, 2111–2112

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 49

direct examination, 2115–2117 endophthalmitis, 1949 epidemiology and transmission, 2113 evaluation, interpretation, and reporting of results, 2123 identification, 2118 isolation, 2117–2118 media, 1959, 1962 microscopy, 1966, 1976, 2111–2112, 2116 nucleic acid detection, 2117 serologic tests, 2120 specimen collection, transport, and processing, 1948–1949, 1951, 2115 staining, 1958 taxonomy, 2110 typing systems, 2119–2120 Blastomyces gilchristii, 2110, 2120 Blastomycosis, 2109–2123 antigen detection, 2116 clinical significance, 2114 collection, transport, and storage of specimens, 1947, 2115 description of agent, 2111 epidemiology and transmission, 2113 evaluation, interpretation, and reporting of results, 2123 nucleic acid detection, 2117 serologic tests, 2120 typing, 2119 Blastoschizomyces, 2001 clinical significance, 1992 description of agents, 1986 epidemiology and transmission, 1992 Blastoschizomyces capitatus, 1984, 1986–1987, 1989, 1998, 2000, 2005 Blastoschizomyces pseudotrichosporon, 1984 Blastospore, 1941 Blattella germanica, 2513 Blautia coccoides, 910 Blautia producta, 909–910, 912 Blautia wexlerae, 910 Bleomycin, 1763 Blepharitis Bacillus oleronius, 443 Demodex, 2330 herpes simplex virus (HSV), 1689 Blood agar, 329 Blood culture automated system, 293 blood volume recommendations, 293 Brucella, 866 critical factors in pathogen recovery, 16–17 agitation of culture bottles, 17 anticoagulant, 17 dilution of blood, 17 medium and additives, 17 number of cultures, 16–17 volume of blood cultured, 16 culture-based detection methods, 19–22 automated systems, 20–22 BacT/Alert, 21, 22 Bactec 9000 series, 21, 22 lysis-centrifugation system, 20 manual blood culture systems, 20 pediatric blood culture bottles, 22 VersaTREK, 21–22, 23 direct rapid antimicrobial susceptibility testing from blood culture bottles, 25 fungal, 1945, 1947–1949 interpretation of blood culture results, 18– 19 quantitative, 18–19

n xlix

semiautomated system, 294 specimen collection, 17–18, 292–294 number and timing of cultures, 18 skin disinfection, 17 transport and handling, 292–294 Blood films, see Thick blood films; Thin blood films Blood parasites antigen and DNA detection, 2307 blood collection for, 2304 blood stains, 2306 buffy coat films, 2307, 2336 concentration procedures, 2307, 2336 detection and identification, 2333–2336 direct detection, 2297, 2306–2307 immunochromatographic tests for malaria, 2335–2336 Knott concentration, 2307, 2336 membrane filtration, 2307, 2336 screening methods, 2307 specimen preparation, procedures, and stains, 2297 staining for, 2334–2335 thick blood films, 2306, 2333–2335 thin blood films, 2306, 2333–2335 Blood specimen fungal, 1945, 1947–1949 laboratory detection of bacteremia and fungemia, 15–26 Mycobacterium, 548 parasitology, 2297, 2304, 2306–2307, 2333–2336 antigen and DNA detection, 2307 blood stains, 2306 buffy coat films, 2307, 2336 collection, 2304 concentration procedures, 2307, 2336 detection and identification, 2333–2336 immunochromatographic tests for malaria, 2335–2336 Knott concentration, 2307, 2336 membrane filtration, 2307, 2336 parasites recovered from, 2294 screening methods, 2307 specimen preparation, procedures, and stains, 2297, 2334–2335 staining for, 2334–2335 thick blood films, 2306, 2333–2335 thin blood films, 2306, 2333–2335 specimen collection, 17–18 collection, transport, and storage guidelines, 272 for parasitology, 2304 viral, 1412–1413 Blood spots, see Dried blood spots Blood stains, for parasites, 2306, 2334–2335 Blood-free Campylobacter selectivity agar, 329 Blood-glucose-cysteine agar, 1959 Blowflies, 2513, 2517 BLSOM (Batch-Learning Self-Organizing Map), 246 BMPA, 893 Bocaparvovirus (genus), 1818 Bocavirus (genus), 1398, 1618, 1823 Bocaviruses, 242, 1406–1407, 1433, 1618; see also Human bocavirus Boceprevir, hepatitis C virus, 1601–1602, 1901 Boceprevir resistance, 1901–1902, 1917 Body fluid specimen, see also specific fluids fungi, 1946, 1948, 1950 Mycobacterium, 548 Body louse, 2510–2511

l

n

SUBJECT INDEX

Boeck and Drbohlav’s Locke-egg-serum (LES) medium, 2315 Bokeloh bat lyssavirus, 1633–1634 BOLD database, 2523 Bolivian hemorrhagic fever, 1674 Bombardier beetles, 2521 Bone infection, see also Osteomyelitis Aggregatibacter, 654 anaerobic Gram-negative rods, 972 Finegoldia magna, 910–911, 911 Fusobacterium, 973 Kingella, 655 microsporidia, 2210 non-spore-forming, anaerobic, Gram-positive rods, 923 Treponema, 1061 Bone marrow specimen/aspirate fungi, 1945, 1947–1949 Gram stain and plating medium recommendations, 286 parasitology, 2294, 2297, 2327–2330 specimen collection, transport, and storage, 272, 291 viruses, 1413 Bone marrow suppression human herpesvirus 6 (HHV-6), 1756 human herpesvirus 8 (HHV-8), 1763 viruses, specimens and methods for detection of, 1406 Bone marrow toxicity chloramphenicol, 1193 nitrofurantoin, 1196 Bone specimen biopsy collection and transport, 271, 281 initial sample handling, 285 Book lice, 2510 Boophilus, 2514 Bordetella, 615, 838–845 antimicrobial susceptibilities, 844–845 clinical significance, 841 collection, transport, and storage of specimens, 841–842 description of genus, 838–840 direct examination, 842 epidemiology and transmission, 840 evaluation, interpretation, and reporting of results, 845 identification, 843 isolation procedures, 842–843 serologic tests, 844 taxonomy, 838 typing systems, 843–844 Bordetella ansorpii, 626–627, 838–841, 844 Bordetella avium, 838–841, 844 Bordetella bronchiseptica, 632–633, 838–844 Bordetella hinzii, 630–631, 838–841 Bordetella holmesii, 626–627, 838–843 Bordetella parapertussis, 624–625 antimicrobial susceptibilities, 844 Bordet-Gengou agar for, 329 clinical significance, 841 collection, transport, and storage of specimens, 300, 841–842 description of, 838–839 direct detection methods, 842 epidemiology and transmission, 840 evaluation, interpretation, and reporting of results, 845 identification, 843 isolation procedures, 842 serologic tests, 844 typing systems, 843–844 Bordetella pertussis, 895 antimicrobial susceptibilities, 844, 1183, 1197

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 50

Bordet-Gengou agar for, 329 clinical significance, 841 collection, transport, and storage of specimens, 300, 841–842 description of, 838–839 direct detection methods, 842 epidemiology and transmission, 840 evaluation, interpretation, and reporting of results, 845 identification, 843 isolation procedures, 842 serologic tests, 844 typing systems, 843–844 Bordetella pertussis selective medium, 329 Bordetella pertussis selective medium with charcoal agar base, 329 Bordetella petrii, 630–631, 838–841 Bordetella trematum, 626–627, 838–841 Bordet-Gengou agar, 329 Boric acid, 2515 Bornaviridae (family), 1398, 1400, 1401 Bornavirus (genus), 1398 Bornholm disease, 1540 Boronic acid, AmpC inhibition by, 1300 Borrelia, 1037–1049 antigen detection, 1043 antimicrobial susceptibilities, 1048 antimicrobial therapy recommendations, 1048 arthropod vectors, 1037–1040 characteristics of, 1037–1039 clinical significance, 996–997, 1040–1041 Lyme borreliosis, 1041 relapsing fever, 1040–1041 collection, transport, and storage of specimens, 283, 1041–1043 blood and serum, 1042 cerebrospinal fluid, 1042–1043 skin biopsy, 1043 synovial fluid/synovial biopsy, 1043 description of genus, 1037–1040 direct examination, 1043 epidemiology and transmission, 1040 evaluation, interpretation, and reporting of results, 1048–1049 genomes, 1040 human humoral immune response to Borrelia antigens, 1044–1045 identification, 994, 996–997, 1044 isolation procedures, 1043–1044 microscopy, 1043 nucleic acid detection techniques, 1043 serodiagnosis error sources in, 1049 influence of antimicrobial therapy on, 1049 two-step approach to, 1045 serologic report, 1048 serological tests, 1044–1048 detection of intrathecally produced (CSF) antibodies, 1047–1048 enzyme immunoassay (EIA), 1044–1046 immunoblotting, 1046–1047 immunofluorescence assay (IFA), 1045– 1046 species diversity, 1039–1040 taxonomy, 1037 vaccine, 1048 Vincent’s angina, 300 Borrelia afzelii, 996, 1038–1040, 1046–1047 Borrelia andersonii, 1038 Borrelia anserina, 1037–1038, 1040 Borrelia bavariensis, 1038–1039

Borrelia bissettii, 1038–1039 Borrelia burgdorferi, 996, 1037–1049 antigen detection, 1043 antimicrobial susceptibilities, 1048, 1183– 1184, 1187 antimicrobial therapy recommendations, 1048 arthropod vectors, 1037–1040, 2507 blood and serum samples, 1042 cerebrospinal fluid sample, 1042–1043 characteristics of, 1038 clinical significance, 1041 collection, transport, and storage of specimens, 283, 1042–1043 detection of intrathecally produced (CSF) antibodies, 1047–1048 direct examination, 1043 epidemiology and transmission, 1040 evaluation, interpretation, and reporting of results, 1048–1049 genomes, 1040 human humoral immune response to Borrelia antigens, 1044–1045 identification, 1044 immunoblotting, 1046–1047 isolation procedures, 1043–1044 microscopy, 1042–1043 nucleic acid detection techniques, 1043 plasmids, 1040 scanning electron micrograph, 1039 serodiagnosis, 1045 serologic report, 1048 serological tests, 1044–1048 skin biopsy specimens, 1043 synovial fluid/synovial biopsy specimens, 1043 Borrelia californiensis, 1038 Borrelia carolinensis, 1038 Borrelia caucasica, 1038 Borrelia coriaceae, 1038, 1040 Borrelia crocidurae, 1038, 1040 Borrelia dipodilli, 1038 Borrelia duttonii, 1037–1038, 1041 Borrelia garinii, 996, 1038–1040, 1046–1047 Borrelia hermsii, 996, 1038–1039, 1042, 1048 Borrelia hispanica, 1038 Borrelia japonica, 1038 Borrelia lonestari, 1038–1039 Borrelia lusitaniae, 1038–1039 Borrelia mazzottii, 1038 Borrelia merionesi, 1038 Borrelia microti, 1038 Borrelia miyamotoi, 1038–1039 Borrelia parkeri, 1038–1039 Borrelia persica, 1038 Borrelia recurrentis, 996, 1037–1038, 1040, 2507 Borrelia sinica, 1038 Borrelia spielmanii, 1038–1039 Borrelia tanukii, 1038 Borrelia theileri, 1038 Borrelia turdi, 1038 Borrelia turicatae, 1038–1039, 1048 Borrelia valaisiana, 1038–1039 Borrelia venezuelensis, 1038 BORSA (borderline oxacillin-resistant S. aureus), 1291–1292 Bortezomib, 1740, 1763 Botflies, 2517–2519 Botryosphaeria theobromae, 2154 Botryosphaeriales (order), 2153–2154, 2162–2163 Botulism, 946–947 biothreat agent, 221

SUBJECT INDEX direct toxin detection, 952 infant, 947 toxin as bioterrorism agent, 947 transmission and disease, 221 wound, 947 Boutonneuse fever, 1124–1126, 1129, 2507, 2513 Bovine albumin fraction V, 320 Bovine albumin Tween 80 medium, 329 Bovine Cryptosporidium dipstick, 2441 Bovine papular stomatitis, 1830 Bovine spongiform encephalopathy (BSE), 206–207, 1861, 1863 Bovine venereal campylobacteriosis, 1000 Bowel obstruction Ascaris lumbricoides, 2451 Diphyllobothrium latum, 2472 Taenia saginata, 2473 Bowie-Dick test, 203 Brachiola, 2209 Brachiola vesicularum, 2209 Brachybacterium chemotaxonomic features, 475 description of genus, 478 identification, 484, 495–496 Brachyspira, 996 antimicrobial susceptibilities, 1072 clinical significance, 1061 collection, transport, and storage of specimens, 1063 description of genus, 1057 direct examination, 1063–1064 epidemiology and transmission, 1059 identification of, 1064–1065 intestinal spirochetosis, 1055, 1058–1059, 1061 clinical significance, 1061 collection, transport, and storage of specimens, 1063 epidemiology and transmission, 1059 isolation procedures, 1064 taxonomy, 1055 typing systems, 1065 Brachyspira aalborgi, 996, 1055, 1057, 1059, 1064–1065 Brachyspira hominis, 1055, 1064 Brachyspira hyodysenteriae, 1055 Brachyspira pilosicoli, 996, 1055, 1057, 1059, 1064–1065, 1072 Bradyrhizobium enterica, 245 Bradyzoites, Toxoplasma gondii, 2373–2374 Brain abscess/infection anaerobic Gram-negative rods, 972 Bacillus licheniformis, 442 Bacteroides, 970 Burkholderia, 794 Citrobacter, 720 Dialister, 974 Fusarium, 2058 Gemella, 424 Gram-positive anaerobic cocci (GPAC), 910–911 Histoplasma capsulatum, 2114 hyaline fungi, 2075–2076 Leuconostoc, 424 Microascus, 2075 Mycoplasma, 1093 Nocardia abscessus, 516 Nocardia cyriacigeorgica, 516 Nocardia farcinica, 517 Nocardia otitidiscaviarum, 517 Nocardia transvalensis, 518 Nocardia veterana, 518 non-spore-forming, anaerobic, Gram-positive rods, 923

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 51

Paenibacillus macerans, 443 Peptostreptococcus anaerobius, 911 phaeohyphomycoses, 2161, 2163 Pseudozyma, 1994 Scopulariopsis, 2075 Serratia, 720 Brain heart infusion, 329 Brain heart infusion agar, 329, 1287–1290, 1295 Brain heart infusion agar (fungal formulation), 1959 Brain heart infusion broth, 1288 Brain tissue specimen fungi, 1946, 1951 parasitology, 2327–2328 rabies diagnosis, 1636 Branched-DNA (bDNA) assays, 55–56, 1443 Branchiibius, 354 Brazilian purpuric fever, 670 Breakpoint susceptibility tests, 1263 Breast abscess anaerobic Gram-negative rods, 974 Brevibacillus parabrevis, 443 Campylobacter, 1001 Corynebacterium confusum, 487 Corynebacterium kroppenstedtii, 490 Finegoldia magna, 911 Prevotella, 972, 973 Breast implant-associated infection Abiotrophia and Granulicatella, 424 Bacillus subtilis, 442 Brevibacillus clinical significance, 443 description of genus, 441–442 identification, 451 taxonomy, 441 Brevibacillus agri, 442 Brevibacillus brevis, 442, 449, 452 Brevibacillus centrosporus, 442–443 Brevibacillus laterosporus, 442, 449, 452 Brevibacillus parabrevis, 442–443 Brevibacterium antimicrobial susceptibility testing, 1328 chemotaxonomic features, 475 clinical significance, 479 description of genus, 475 epidemiology and transmission, 478 identification, 438, 494 Brevibacterium casei, 494 Brevibacterium helvolum, see Pseudoclavibacter Brevibacterium luteolum, 494 Brevibacterium mcbrellneri, 494 Brevibacterium sanguinis, 494 Brevibacterium stationis, 492; see also Corynebacterium stationis Brevundimonas, 615 characteristics of, 802 clinical significance, 795 description of genus, 792 epidemiology and transmission, 793 identification, 802 taxonomy, 792 Brevundimonas diminuta, 626–629, 792, 795, 802 Brevundimonas vancanneytii, 792 Brevundimonas vesicularis, 628–629, 792, 795–796, 802 Bright-field microscopy, see also Microscopy compound microscope, 7–9 typical configuration for, 6 Brilliance Candida agar, 1999 Brilliance CRE agar, 329 Brilliance ESBL agar, 329 Brilliance green agar, 330

n li

Brilliance green-phenol red agar, 330 Brilliance MRSA agar, 329 Brilliance Salmonella agar, 329–330 Brilliance Staph 24 agar, 330 Brilliance UTI agar, 330 Brilliance UTI clarity agar, 330 Brilliance VRE agar, 330 Brill-Zinsser disease, 1125 Brincidofovir, 1720, 1886 British Society for Antimicrobial Chemotherapy, 1253, 1268–1269, 1297 Brivudine, for varicella-zoster virus, 1706 Bromcresol purple (BCP)-milk solids-glucose (BCPMSG), 2141 Bromcresol purple-deoxycholate agar, 330 Bromcresol purple-milk solids-glucose medium, 1959 Bronchial aspirate specimen fungi, 1946, 1950 Mycobacterium, 547 Bronchiectasis Mycobacterium avium complex, 541, 599 Mycobacterium chelonae/M. abscessus group, 599 Mycobacterium xenopi, 543 Pasteurella, 655 Pseudomonas aeruginosa, 775 Bronchiolitis adenoviruses, 1771 coronaviruses, 1569 enterovirus, 1540 human metapneumovirus, 1509 microsporidia, 2210 respiratory syncytial virus (RSV), 1500 viruses, specimens and methods for detection of, 1407 Bronchiolitis obliterans, adenoviruses and, 1773 Bronchitis adenoviruses, 1771, 1772 Chlamydia pneumoniae, 1108 enterovirus, 1540 etiologies, usual, 290 Haemophilus influenzae, 669 Haemophilus parainfluenzae, 670 microsporidia, 2213 Nocardia cyriacigeorgica, 516 Paragonimus, 2487 Bronchoalveolar brush specimens Gram stain and plating medium recommendations, 286 specimen collection, transport, and handling, 298 Bronchoalveolar lavage (BAL) specimen Aspergillus, 1971, 2037–2039 coronaviruses, 1569–1570 fungi, 1946, 1948, 1950 Gram stain and plating medium recommendations, 286 Mycobacterium, 547 Pneumocystis, 2020 respiratory syncytial virus, 1501 screening specimens, 284 specimen collection, transport, and handling, 278, 298, 1415 varicella-zoster virus detection, 1710 viral infections, 1415 Bronchoscopy aspirates for parasitology, 2331 specimen collection, transport, and handling, 298 Broth macrodilution method antifungal susceptibility testing

lii

n

SUBJECT INDEX

Broth macrodilution method (continued) molds, 2268, 2270 yeasts, 2262 antimicrobial susceptibility testing, 1258, 1261–1262 advantages and disadvantages, 1262 dilution of antimicrobial agents, 1258 incubation, 1261 inoculation procedures, 1261 interpretation and reporting of result s, 1261–1262 preparation, supplementation, and storage of media, 1258, 1261 Broth microdilution antimicrobial susceptibility testing, 1262–1263 ability to detect resistance, 1278–1280 aminoglycoside resistance in Enterococcus, 1278 carbapenem resistance, 1279–1280 ESBL-producing Enterobacteriaceae, 1279 glycopeptide susceptibility reduction in staphylococci, 1279 in Gram-negative bacteria, 1280 in Gram-positive bacteria, 1279 inducible clindamycin resistance, 1279 linezolid resistance in enterococci and staphylococci, 1278 oxacillin resistance in staphylococci, 1278 penicillin resistance in staphylococci, 1278 Streptococcus resistance, 1279 vancomycin resistance in enterococci, 1278 advantages and disadvantages, 1263 antifungal susceptibility testing dermatophytes, 2271 molds, 2268–2271 yeasts, 2258–2264 automated, 1275–1280 ability to detect resistance, 1278–1280 advantages, 1277–1278 BD Phoenix system, 1277 disadvantages, 1278 MicroScan WalkAway system, 1276– 1277 Sensititre ARIS 2X, 1277 VITEK systems, 1275–1276 breakpoint susceptibility tests, 1263 dilution of antimicrobial agents, 1262 for aminoglycoside resistance detection in enterococci, 1288 for anaerobic bacteria, 1344–1345 commercial panels, 1345 incubation conditions, 1344 inoculation procedure, 1344 inoculum preparation, 1344 interpretation and result reporting, 1345 medium, 1344 quality control, 1344–1345 for extended-spectrum β-lactamase (ESBL) production confirmation, 1266 fastidious bacteria, 1315, 1317–1318 gradient diffusion method, 1263 incubation, 1262–1263 inoculation procedures, 1262 interpretation and reporting of results, 1263 manual, 1275 metallo-β-lactamase detection, 1301 potential agents of bioterrorism, 1316 preparation, supplementation, and storage of media, 1262 resistance screens, 1263

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 52

semiautomated, 1275 Brown dog tick, 2513 Brown recluse spider, 2521 Browntail moth, 2518 Brucella, 222, 824, 863–869 antibiotic resistance, 1234 antigenic components, 863 antimicrobial susceptibilities, 868, 1179, 1186, 1192, 1195 antimicrobial susceptibility testing, 1316, 1324–1325 automated identification, 32 biothreat agent, 220, 222 blood culture, 18 characteristics, 220 clinical features, 864–865 collection, handling, storage, and transport of specimens, 865 complications, 865 culture, 865–866 description of genus, 863 differentiation of Francisella from, 852 direct detection, 865 epidemiology and transmission, 222, 864 evaluation, interpretation, and reporting of results, 869 identification, 866–867 immune response, 864 laboratory-acquired infections, 176, 864 pathogenic mechanisms, 864 prevention, 869 serologic tests, 867–869 taxonomy, 863 treatment, 868–869 typing systems, 867 vaccine, 869 virulence factors, 864 Brucella abortus, 863–864, 868–869 Brucella agar, 330, 1342, 1344 Brucella agar base campylobacter medium, 330 Brucella agar base with blood and selective supplement, 330 Brucella blood culture broth, 330–331 Brucella broth, 331, 1344 Brucella canis, 863–864, 866, 868 Brucella ceti, 863 Brucella delphini, 863 Brucella inopinata, 863 Brucella laked sheep blood agar with kanamycin and vancomycin, 331 Brucella melitensis, 863–864, 866–869, 1180 Brucella neotomae, 863–864 Brucella ovis, 863–864 Brucella pinnipedialis, 863 Brucella suis, 863–864, 869 Brucellacapt, 867–868 Brucellaceae (family), 824, 863 Brucella-containing vacuoles, 864 Brucellosis, 863–865 biothreat agent, 222 laboratory tests suggested for, 125 laboratory-acquired infections, 176 transmission and disease, 222 Brugia, 2461–2465 arthropod vector, 2507 clinical significance, 2462, 2464 description of agent, 2461–2462, 2464 detection, 2328, 2332 diagnosis, 2464 direct examination, 2464–2465 epidemiology and transmission, 2462 microscopy, 2464–2465 nucleic acid detection, 2465

prevention, 2465 serologic tests, 2465 taxonomy, 2461 treatment, 2465, 2534–2535 Brugia malayi, 1139, 2461–2465 clinical significance, 2462, 2464 description of agent, 2461–2462, 2464 detection, 2328, 2332 direct examination, 2464–2465 epidemiology and transmission, 2462 microscopy, 2464–2465 prevention, 2465 serologic tests, 2465 taxonomy, 2461 treatment, 2465, 2531, 2534–2535 Brugia timori, 2461–2462, 2465, 2535 Bruker MALDI Biotyper, see Biotyper system Bryantella formatexigens, see Marvinbryantia formatexigens BSE (bovine spongiform encephalopathy), 206–207, 1861, 1863 BsoB1 enzyme, 64 BTA-798, 1558 Bubonic plague, 741 Budding, 1935, 1938, 1939 Budvicia aquatica, 718, 727 Buerger’s disease, 971 Buffalopox virus, 1828–1829 Buffered charcoal-yeast extract (BCYE) agar, 2315 Buffered charcoal-yeast extract agar with cysteine (BCYE alpha base), 331 Buffered charcoal-yeast extract differential agar, 331 Buffered formalin, 2310 Buffers, 320 Buffy coat films, for parasites, 2307, 2336 Bulleidia, 921 Bulleidia extructa, 922, 924, 930 Bull’s eye rash, 1041 Bumblebees, 2518 Bundibugyo virus, 1670, 1672–1673, 1682 Bunyamwera virus, 1645 Bunyaviridae (family), 1399–1401, 1644–1645, 1660, 1669 Bunyavirus (genus), 1644, 1647–1648, 1651–1653, 1655 Burkholderia, 614, 774 antimicrobial susceptibilities, 803–804, 1175, 1180 classification in genus, 264 clinical significance, 793–794 description of genus, 792 direct examination, 795–796 epidemiology and transmission, 792–793 evaluation, interpretation, and reporting of results, 804 identification, 797–800 isolation procedures, 796–797 serologic tests, 803 taxonomy, 791 typing systems, 803 Burkholderia ambifaria, 792–793, 798 Burkholderia anthina, 792–793, 798 Burkholderia arboris, 793, 798 Burkholderia cenocepacia, 630–633, 791–793, 798 Burkholderia cepacia, 615, 626–627, 798, 1192–1193 antimicrobial susceptibility testing, 1255–1256, 1266, 1270 in cystic fibrosis patients, 299 specimen collection, transport, and handling, 299

SUBJECT INDEX Burkholderia cepacia agar, 331 Burkholderia cepacia complex antimicrobial susceptibilities, 803, 1176–1177 characteristics of, 798 clinical significance, 793–794 direct examination, 795 epidemiology and transmission, 792 evaluation, interpretation, and reporting of results, 804 identification, 797–800 isolation procedures, 797 ribotyping, 803 taxonomy, 791 Burkholderia cepacia selective agar, 331 Burkholderia cocovenenans, 791 Burkholderia contaminans, 793, 798 Burkholderia diffusa, 793, 798 Burkholderia dolosa, 793, 798–799 Burkholderia endofungorum, 791 Burkholderia fungorum, 791 Burkholderia gladioli, 626–627 characteristics of, 798 clinical significance, 793–794 epidemiology and transmission, 793 identification, 797–799 taxonomy, 791 Burkholderia glumae, 791 Burkholderia insidiosa, 791 Burkholderia lata, 793, 798 Burkholderia latens, 793, 798 Burkholderia mallei antimicrobial susceptibilities, 804 antimicrobial susceptibility testing, 1316, 1324–1325 biothreat agent, 220, 222 characteristics, 220, 800 clinical significance, 794 description, 792 epidemiology and transmission, 793 evaluation, interpretation, and reporting of results, 804 identification, 799–800 taxonomy, 791 transmission and disease, 222 Burkholderia mannitolilytica, 791 Burkholderia metallidurans, 793 Burkholderia multivorans, 630–631, 791–793, 798–799 Burkholderia pickettii, 791 Burkholderia pseudomallei, 630–631, 778 antimicrobial susceptibilities, 803–804, 1186, 1192 antimicrobial susceptibility testing, 1316, 1324–1325 Ashdown medium for, 328 β-lactamase, 1325 biothreat agent, 220, 222 characteristics, 220, 800 clinical significance, 794 collection of specimens, 795 colony morphology, 797 direct examination, 795–796 epidemiology and transmission, 792–793 evaluation, interpretation, and reporting of results, 804 identification, 799–800 in Acanthamoeba, 2389 isolation procedures, 796–797 laboratory-acquired infections, 176–177 ribotyping, 803 serologic tests, 803 taxonomy, 791 transmission and disease, 222

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 53

Burkholderia pseudomallei selective agar, 331 Burkholderia pseudomallei selective agar (BPSA), 797 Burkholderia pseudomultivorans, 793, 798–799 Burkholderia pyrrocinia, 792–793, 798 Burkholderia rhizoxinica, 791 Burkholderia seminalis, 793, 798 Burkholderia stabilis, 793, 798 Burkholderia thailandensis, 791, 799–800 Burkholderia tropica, 791 Burkholderia ubonensis, 794, 798 Burkholderia vietnamiensis, 793, 798–799 Burkholderiales (order), 838 Burkitt’s lymphoma, 1738–1740 Burns Bacteroides, 971 Fusarium, 2058 Pseudomonas aeruginosa, 775 specimen collection, transport, and storage guidelines, 273 Bursitis Mycobacterium szulgai, 543 Stenotrophomonas maltophilia, 794 Buruli ulcer, 543 Bush-Jacoby classification system for β-lactamases, 1223 Bussuquara virus, 1645 Buthacus, 2520 Buthus, 2520 Butterflies, 2518–2519, 2522 Buttiauxella gaviniae, 718 Buttiauxella noackiae, 718 Bwamba virus, 1645 C. albicans PNA FISH assay, 1997 C3d receptor, Epstein-Barr virus and, 1738 Cabinets, biosafety, 172–173, 282–283 Cadophora, 2158 Calabar swelling, 2467 Calamine, 2515 Calcofluor white, 1957 Calcofluor white stain, 1970, 1974, 1975, 2316 Calibrated dichotomous sensitivity (CDS) disk diffusion method, 1268–1269 CaliciNet Network, 151 Caliciviridae (family), 1399–1401, 1617–1618 Caliciviruses antigen detection, 1623–1625 cell culture, 1627 clinical significance, 1620–1622 description of agents, 1618–1619 epidemiology and transmission, 1620–1621 molecular detection assays, 1625–1627 serologic tests, 1628 specimen collection and handling, 1406 taxonomy, 1617–1618 Calidifontibacter, 354 Calliphora, 2519 Calliphoridae (family), 2513 Calosphaeriales (order), 2153–2155, 2163–2164 Camera systems, for photomicroscopy, 11–12 CAMP test, 317 coryneform Gram-positive rods, 481, 487, 489–494, 496 Listeria, 464 Staphylococcus aureus, 392, 481 Streptococcus agalactiae, 392 Campy-CVA, 1002–1003 Campy-JCL, 1005 Campylobacter, 974, 998–1007 antigen detection, 1002

n liii

antimicrobial susceptibilities, 1006–1007, 1180, 1183 antimicrobial susceptibility testing, 1326–1327 clinical significance, 996–997, 1000–1001 collection, transport, and storage of specimens, 1001 description of agents, 998 direct examination, 1001–1002 epidemiology and transmission, 998–1000 evaluation, interpretation, and reporting of results, 1007 Helicobacter and, 1013–1014, 1021 identification, 994–997, 1003–1006 isolation procedures, 1002–1003 microscopy, 1001–1002 molecular serotyping, 145 nucleic acid amplification tests (NAATs), 1002 reservoirs, 999 serologic tests, 1006 Skirrow brucella medium, 344 subtyping, 139 taxonomy, 998 typing systems, 1006 Campylobacter agar, Blaser’s, 331 Campylobacter agar, Skirrow’s, 331 Campylobacter avium, 999, 1001, 1004 Campylobacter blood agar, 331 Campylobacter canadensis, 999, 1001, 1004 Campylobacter charcoal differential agar, 331 Campylobacter coli, 996, 1021 antigen detection, 1002 antimicrobial susceptibilities, 1006–1007 antimicrobial susceptibility testing, 1317, 1326–1327 evaluation, interpretation, and reporting of results, 1007 identification, 1004–1006 isolation procedures, 1002–1003 reservoirs, 999 source attribution, 147 specimen collection, transport, and storage guidelines, 301–302, 1001 typing, 1006 Campylobacter concisus, 996–1002, 1004 Campylobacter cuniculorum, 999, 1001, 1004 Campylobacter curvus, 996–999, 1001–1002, 1004 Campylobacter fetus, 1001, 1006 Campylobacter fetus subsp. fetus, 996–997, 999–1000, 1004–1005, 1007 Campylobacter fetus subsp. venerealis, 999–1000, 1004 Campylobacter gracilis, 997, 999, 1001, 1004, 1348 Campylobacter helveticus, 999, 1001, 1004, 1006 Campylobacter hominis, 996, 998–999, 1001, 1004 Campylobacter hyointestinalis, 996, 1000, 1002, 1006 Campylobacter hyointestinalis subsp. hyointestinalis, 999, 1004 Campylobacter hyointestinalis subsp. lawsonii, 999, 1004 Campylobacter insulaenigrae, 996, 999, 1004 Campylobacter jejuni, 1014, 1021 antigen detection, 1002 antimicrobial susceptibilities, 1006–1007, 1180, 1195 antimicrobial susceptibility testing, 1317, 1326–1327

liv

n

SUBJECT INDEX

Campylobacter jejuni (continued) clinical significance, 1000 epidemiology, 998–999 evaluation, interpretation, and reporting of results, 1007 identification, 1004–1006 isolation procedures, 1002–1003 source attribution, 147 specimen collection, transport, and storage guidelines, 301–302, 1001 typing, 1006 Campylobacter jejuni subsp. doylei, 996, 999–1000, 1004–1005 Campylobacter jejuni subsp. jejuni, 996–998, 1004 Campylobacter lanienae, 996, 999, 1001, 1004, 1006 Campylobacter lari, 1000, 1005–1007 Campylobacter lari subsp. concheus, 996–1000, 1004 Campylobacter lari subsp. lari, 996–1000, 1004 Campylobacter mucosalis, 999, 1001–1002, 1004 Campylobacter peloridis, 996, 999, 1001, 1004 Campylobacter pyloridis, see Helicobacter pylori Campylobacter rectus, 996–999, 1001–1002, 1004 Campylobacter selective medium, 331 Campylobacter showae, 997–999, 1001, 1004 Campylobacter sputorum, 996, 1001–1002 Campylobacter sputorum bv. faecalis, 999, 1004 Campylobacter sputorum bv. paraureolyticus, 999, 1001, 1004 Campylobacter sputorum bv. sputorum, 999, 1004 Campylobacter subantarcticus, 998–999, 1004 Campylobacter thioglycolate (Campy-Thio) medium, 331 Campylobacter upsaliensis, 996–997, 999–1007 Campylobacter ureolyticus, 967, 997–999, 1004 Campylobacter volucris, 998–999 Campylobacteraceae (family), 998–1000 Canavanine-glycine-bromthymol blue agar, 1959–1960 Candida, 2115, 2139 antibody detection, 1969, 1971 antifungal resistance, 2225–2227, 2229, 2237–2246 antifungal susceptibilities, 1183, 2004–2006, 2224 antifungal susceptibility testing, 2255–2267, 2273 antigen detection, 1972, 1975, 1977, 1996 antimicrobial susceptibility testing, 1246, 2255–2268 biofilms, 2242–2243 clinical significance, 1993 cycloheximide inhibition, 1951, 1955 description of agents, 1986–1988 detection, 2327 endocarditis, 1948 endophthalmitis, 1949 epidemiology and transmission, 1992 evaluation, interpretation, and reporting of results, 2006 identification, 1978, 1998–2004 media, 1952–1953, 1959–1961 microscopy, 1966, 1998 nucleic acid detection, 1979, 1997–1998 serologic tests, 2004 specimen collection, transport, and processing, 1944, 1947–1953

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 54

taxonomy, 1984–1985 Candida aaseri, 1985, 1993 Candida africana, 1984, 1986, 1992 Candida africana var. africana, 1986 Candida albicans, 2139, 2415 antifungal resistance, 2225–2226, 2229, 2237–2246 antifungal susceptibility, 2005, 2224 antifungal susceptibility testing, 2256, 2262, 2264–2265, 2267 antigen detection, 1975, 1996 biofilms, 2242–2243 blood culture, 18 chromogenic agar, 1999 clinical significance, 1993 cultural and biochemical characteristics, 1989 description, 1986–1988 endocarditis, 1948 epidemiology and transmission, 1991–1992 germ tube test, 1999 identification, 1978, 1998–1999, 2001– 2003 isolation, 1998 keratitis, 1949 media, 1959–1960 microscopy, 1973, 1998 multilocus sequence typing (MLST), 2004 nucleic acid detection, 1997, 2415 specimen collection, transport, and processing, 1950 taxonomy, 1984–1985 typing, 2004 Candida auris, 1985, 1993 Candida bracarensis, 1985, 1993 Candida catenulata, 1989, 1993 Candida chromogenic agar, 1960 Candida ciferrii, 1985, 1993, 2000 Candida claussenii, 1984 Candida diagnostic agar (CDA), 1960 Candida dubliniensis, 1985–1986, 1989, 1991, 1993, 1997–1999, 2002–2003, 2005, 2243 Candida famata, 1985–1986, 1989, 1993, 2000, 2005 Candida famata var. flareri, 1985–1986 Candida fermentati, 1985 Candida glabrata, 1985, 1989, 1991–1993, 1997–1998 antifungal resistance, 2006, 2225, 2227, 2229, 2237–2238, 2241, 2243– 2246 antifungal susceptibility, 2004–2006, 2224 antifungal susceptibility testing, 2256– 2257, 2263–2265, 2267 antigen detection, 1975 clinical significance, 1993 cultural and biochemical characteristics, 1989 description, 1986 epidemiology and transmission, 1992 identification, 2001–2004 media, 1960 microscopy, 1973 multilocus sequence typing (MLST), 2004 nucleic acid detection, 1979 Rapid Trehalose test, 2001 Candida guilliermondii, 1985, 1989, 1993, 2005, 2229, 2237–2238, 2245, 2264 cultural and biochemical characteristics, 1989 description, 1986–1987 identification, 2000–2001, 2003 Candida guilliermondii var. membranifaciens, 1985

Candida haemulonii, 1985, 1993, 2005 Candida inconspicua, 1993, 2001, 2004–2005 Candida kefyr, 1985, 1989, 1993, 2000–2001, 2004–2005 Candida krusei, 1985–1987, 1992–1993 antifungal resistance, 2006, 2238, 2241, 2243 antifungal susceptibility, 2004–2006, 2224 antifungal susceptibility testing, 2256, 2261, 2264, 2267, 2272 cultural and biochemical characteristics, 1989 description, 1986–1987 identification, 1998–2003 media, 1960 Candida lambica, 1989 Candida langeronii, 1984 Candida lipolytica, 1985–1986, 1989–1990, 1993, 2000–2001 Candida lusitaniae, 1985–1989, 1991, 1993, 1997 antifungal resistance, 2229, 2238, 2243, 2246 antifungal susceptibility, 2004–2005, 2224 description, 1986–1988 identification, 2000–2001, 2003 Candida metapsilosis, 1984–1985, 2003, 2237, 2245 Candida neorugosa, 1985 Candida nivariensis, 1985–1986, 1993 Candida norvegensis, 1985, 1993, 2000, 2004 Candida orthopsilosis, 1984, 1997, 2003 Candida parapsilosis, 1984–1985, 1991–1993, 1997, 2001, 2003, 2139 antifungal resistance, 2006, 2237–2238, 2243, 2245 antifungal susceptibility, 2005, 2224 antifungal susceptibility testing, 2256, 2261–2262, 2264 cultural and biochemical characteristics, 1989 keratitis, 1949 morphologic features, 1987 Candida pararugosa, 1985 Candida pelliculosa, 1988–1989, 1993, 2000, 2238 Candida pintolopesii, 1989, 2000 Candida pseudoaaseri, 1985 Candida pseudorugosa, 1985 Candida pulcherrima, 1993, 2000 Candida rugosa, 1985, 1989, 1993, 2005, 2243 Candida stellatoidea, 1984–1985 Candida subhashii, 1985, 1993 Candida tropicalis, 1991–1993, 1997–1998, 2006 antifungal resistance, 2229, 2243 antifungal susceptibility, 2005, 2224 antifungal susceptibility testing, 2256 antigen detection, 1975 clinical significance, 1993 cultural and biochemical characteristics, 1989 description, 1986–1987 epidemiology and transmission, 1992 identification, 1998–2003 keratitis, 1949 media, 1959 Candida utilis, 1985, 1993, 2000 Candida viswanathii, 1993 Candida zeylanoides, 1985, 1989, 1991, 1993 “Candidatus,” 259 Candidemia, 1992 Candidiasis, 1993, 2415

SUBJECT INDEX mucocutaneous, 1993 specimens for, 1947, 1949–1951 vaginal, 1951 CandiSelect 4, 1960, 1999 Cand-Tec latex agglutination test, 1972, 1996 Canine granulocytotropic ehrlichiosis, 1139 Canonical pathogens, 227 Cantharidin, 2521 Capillaria, 2494, 2497, 2531 Capillaria hepatica, 2328, 2330 Capillaria philippinensis, 2493, 2497 clinical significance, 2497 description of agent, 2497 direct examination by microscopy, 2497 epidemiology, transmission, and prevention, 2497 serologic tests, 2497 treatment, 2497 Capillariasis, 2497 Capita Syphilis-G, 1067, 1071 Capita Syphilis-M, 1067, 1071 Capnocytophaga antimicrobial susceptibilities, 662 clinical significance, 654 direct examination, 656 epidemiology and transmission, 653–654 identification, 658–659 isolation procedures, 656 serotyping, 661 taxonomy and description of, 652–653 Capnocytophaga canimorsus, 652–654, 656, 659 Capnocytophaga cynodegmi, 652–654, 659 Capnocytophaga gingivalis, 652–654, 659 Capnocytophaga granulosa, 652–654, 659 Capnocytophaga haemolytica, 652–654, 659 Capnocytophaga leadbetteri, 652–654 Capnocytophaga ochracea, 652–654, 657, 659 Capnocytophaga sputigena, 652–654, 659 Capnodiales, 2162 Capreomycin, for Mycobacterium infection, 1358, 1360 Capreomycin resistance, 1356 Capronia semiimmersa, 2154 Captia HSV1 and HSV2 IgG ELISA kits, 1693 Capture assays, IgM, 100 Carba NP test, 1225, 1301 Carbapenem(s), 1172, 1176–1177 adverse effects, 1177 antimicrobial susceptibility testing, 1255, 1260 mechanism of action, 1176 pharmacology, 1176 spectrum of activity, 1177 structure, 1172 Carbapenem resistance, 1176–1177 Bacillus cereus, 1326 Bacteroides fragilis group, 1346–1347 detection by automated antimicrobial susceptibility testing, 1279–1280 Carbapenemases, 1176, 1178, 1223–1226 Enterobacteriaceae, 728, 730, 1300–1302 Pseudomonas aeruginosa, 781–782, 1300–1301 tests for detection of, 1300–1302, 1383 Carbapenem-resistant Enterobacteriaceae commercial sources of chromogenic agar media for, 327 media for detection, 324 Carbarsone, for Dientamoeba fragilis, 2413 Carbohydrate assimilation tests, yeasts, 2001 Carbohydrate fermentation tests, for yeast identification, 2001

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 55

Carbohydrate utilization aerobic Gram-negative bacteria identification, 614 Clostridium, 954–955 Mycobacterium, 601 Neisseria, 640–641 Carboxypenicillins, 1173 Card agglutination trypanosomiasis test (CATT), 2367 Cardiac arrhythmias, Leptospira and, 1030 Cardiac disease, see also Cardiomyopathy; Cardiovascular disease Brucella, 865 Epstein-Barr virus, 1739 respiratory syncytial virus (RSV), 1500 viruses, specimens and methods for detection of, 1406 Cardiac dysrhythmias, macrolides and, 1183 Cardiobacteriaceae (family), 652–653 Cardiobacterium antimicrobial susceptibilities, 662 antimicrobial susceptibility testing, 1328 clinical significance, 654 direct examination, 656 epidemiology and transmission, 653 identification, 658, 660 isolation procedures, 656 taxonomy and description of, 653 Cardiobacterium hominis, 653–654, 656, 658, 662, 1328 Cardiobacterium valvarum, 653–654, 658, 662 Cardiomyopathy human herpesvirus 6 (HHV-6), 1756 Loa loa, 2467 Trypanosoma cruzi, 2363, 2365 Cardiovascular disease rubella, 1526 syphilis, 1059 Cardiovirus (genus), 1551 Carditis, Borrelia, 1041 CARDS (community-acquired respiratory distress syndrome) toxin, 1088–1089, 1094 CareHPV test, 1790 Carnobacteriaceae (family), 354, 356 Carpet beetles, 2521 Carrion’s disease, 873, 876 Cary-Blair transport medium, 331–332 CAS virus, 1672 CA-SFM (Société Française de Microbiologie), 1268–1269 Caspofungin, 2228 antifungal susceptibility testing, 2255–2273 Aspergillus, 2044 Candida, 2004–2005 dimorphic fungi, 2122 eumycotic mycetoma fungi, 2181–2182 spectrum of activity, 2224 yeast species, MICs for, 2005 Caspofungin resistance, 2239, 2243 Castaneda bottle, 20 Castor bean tick, 2512 Cat scratch disease, 283, 873–874, 876, 882 Catabacter, 921 Catabacter hongkongensis, 922 Catalase test, 317 Catalase-negative, Gram-positive cocci, 422–431; see also specific genera antimicrobial susceptibilities, 429–430 with cells arranged in clusters or irregular groups, 427 with cells arranged in pairs or chains, 426 cellular morphology, 423

n lv

clinical significance, 424–425 collection, transport, and storage of specimens, 425 description of genera, 423 direct examination, 425 epidemiology and transmissions, 423–424 evaluation, interpretation, and reporting of results, 430–431 identification, 425–429 isolation procedures, 425 serologic tests, 429 taxonomy, 422–423 typing systems, 429 Catalase-positive Gram-positive cocci, 354–372 Catalogue, strain, 150–151 Cataracts, rubella, 1526 Catarina virus, 1669, 1672 Catenate, 1941 Catenibacterium, 921 Catenibacterium mitsuokai, 920, 930 Caterpillars, urticating, 2518–2519 Catheter specimen collection, transport, and storage guidelines, 273 fungi, 1945, 1949 Catheter-associated infection Acinetobacter, 813 Bacillus licheniformis, 442 Bacillus pumilus, 443 Candida, 1993 Comamonas testosteroni, 795 Corynebacterium tuberculostearicum, 479 Fusarium, 2058 Mycobacterium, 600 Nocardia otitidiscaviarum, 517 Ochrobactrum, 824 Paenibacillus macerans, 443 Pseudomonas, 776 Pseudozyma, 1994 Ralstonia, 795 Rhodotorula, 1994 Rothia mucilaginosa, 361 Serratia, 720 Staphylococcus, 360 Tsukamurella tyrosinosolvens, 519 Cation-adjusted Mueller-Hinton broth (CAMHB), 1258, 1261–1262, 1290, 1296, 1298, 1317–1318, 1321, 1324–1325, 1327–1328, 1330 Cats, in Toxoplasma gondii life cycle, 2373–2375 CATT (card agglutination trypanosomiasis test), 2367 Caudovirales (order), 1402 CCR5 coreceptor, 1437, 1447–1448, 1920 CCR5 inhibitor resistance, HIV, 1897, 1899 ccrB typing tool, 150 CD4+ T-cell, human immunodeficiency virus and, 1439–1440, 1449 CDC, see Centers for Disease Control and Prevention CDC anaerobe 5% sheep blood agar, 332 CDS (calibrated dichotomous sensitivity) disk diffusion method, 1268–1269 Cedecea, 715, 721, 725 Cedecea davisae, 718, 725 Cedecea lapagei, 718, 725 Cedecea neteri, 718 Cefaclor, 1173–1174, 1198 Cefadroxil, 1173, 1198 Cefazolin, 1198, 1255, 1259 Cefdinir, 1173, 1175, 1198 Cefditoren, 1173, 1175, 1198

lvi

n

SUBJECT INDEX

Cefepime, 1175–1176, 1198, 1255, 1259 Cefinase disk method, 1328 Cefixime, 1173, 1175, 1198 Cefoperazone, 1174–1175, 1198 Cefotaxime, 1173, 1174, 1198, 1255, 1259 Cefotetan, 1174, 1175, 1198, 1255, 1259 Cefotetan resistance Bacteroides fragilis group, 1346 Clostridium innocuum, 1348 Cefoxitin, 1174, 1198 anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1255, 1259 as surrogate for oxacillin, 1291 Bacteroides fragilis group susceptibility percentages, 1350 Cefoxitin resistance, 1300, 1346, 1348 Cefpirome, 1198 Cefpodoxime, 1173, 1175, 1198, 1259 Cefprozil, 1173–1174, 1198 Cefsulodin-irgasan-novobiocin agar, 332 Ceftaroline, 1173, 1175, 1198 antimicrobial susceptibility testing, 1255, 1259 with avibactam, 1178 Ceftazidime, 1174–1175, 1198 antimicrobial susceptibility testing, 1255, 1259 with avibactam, 1178 Ceftazidime resistance, 1325 Ceftibuten, 1173, 1175, 1198 Ceftobiprole, 1173, 1175, 1198 Ceftolozane, 1175 Ceftolozane with tazobactam, 1178 Ceftriaxone, 1173–1175, 1198, 1255, 1259 Ceftriaxone resistance, 1322, 1383 Cefuroxime, 1173, 1174, 1198, 1255, 1259 Celera Assembler, 233 Cell culture cytotoxicity neutralization assays (CCCNAs), Clostridium difficile and, 950–951 Cell culture media, 1429–1430 Cell cytotoxicity assays, 1429 Cell membrane analysis, of aerobic actinomycetes, 523 Cell surface protein gene (CSP) typing, for Aspergillus, 2043 Cell wall analysis of aerobic actinomycetes, 523 composition, 262 Mycobacterium, 536–537 Cell-mediated immunity cytomegalovirus (CMV), 17, 22, 1730 Toxoplasma gondii, 2376 varicella-zoster virus, 1712 Cellobiose-arginine-lysine agar, 332 Cellular fatty acids analysis, 262 in Corynebacterium, 474–475, 483 Cellulitis Aeromonas, 754 aspirate specimen collection, transport, and storage guidelines, 273 Clostridium, 946 Erysipelothrix, 468 Gram-positive anaerobic cocci (GPAC), 910 Haemophilus influenzae, 669 hyaline fungi, 2076 Mycobacterium chelonae, 596 Mycobacterium haemophilum, 542 Mycobacterium smegmatis, 598 Nocardia brasiliensis, 516

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 56

non-spore-forming, anaerobic, Grampositive rods, 923 Pseudomonas, 776 sample handling, 292 Vibrio fluvialis, 765 Vibrio vulnificus, 765 Cellulomonadaceae (family), 1159 Cellulomonas antimicrobial susceptibility testing, 1328 clinical significance, 479 description of genus, 477 epidemiology and transmission, 479 identification, 438, 484, 495 taxonomy, 474–475 Cellulomonas cellulans, 495 Cellulomonas denverensis, 477, 495 Cellulomonas fermentans, 477 Cellulomonas hominis, 477, 495 Cellulomonas humilata, 477 Cellulose tape preparations, 2324 Cellulosimicrobium clinical significance, 479 description of genus, 477 epidemiology and transmission, 479 identification, 438, 484, 495 taxonomy, 474–475 Cellulosimicrobium cellulans, 477, 495 Cellulosimicrobium funkei, 477, 495 Centers for Disease Control and Prevention (CDC) bioterrorism agents, 1324 bioterrorism agents and diseases, 217–219 Cholera and Other Vibrio Illness Surveillance (COVIS) system, 769 disinfection and, 193, 196–197, 200–202, 206–207 hand hygiene, 186–187 Helicobacter pylori Antimicrobial Resistance Monitoring Program, 1328 human immunodeficiency virus disease classification, 1440 Human Influenza Virus Real-time RT-PCR Detection and Characterization Panel, 1477 Model Performance Evaluation Program, 1365 outbreak response, 121 tuberculosis, 1362, 1365, 1367–1368 Vaccine Preventable Disease Reference Centers (VPD RCs), 1523 Centers for Medicare and Medicaid Services, 217 Centipeda, 969, 982 Centipeda periodontii, 969, 974 Centipedes, 2520, 2522 Centraalbureau voor Schimmelcultures (CBS) Yeast database, 2002 Central nervous system disease/infection Abiotrophia and Granulicatella, 424 anaerobic Gram-negative rods, 972 Anaplasma phagocyrophilum, 1139 arboviruses, 1647 arenaviruses, 1673 BK polyomavirus, 1805 Borrelia, 1041 Brucella, 865 cytomegalovirus, 1719 Ehrlichia chaffeensis, 1138 etiologies, usual, 290 Gnathostoma, 2498 herpes simplex virus (HSV), 1689 Histoplasma capsulatum, 2114 human herpesvirus 7 (HHV-7), 1761

human immunodeficiency virus, 1439–1440 lymphoma and Epstein-Barr virus, 1747 measles, 1520 Mycobacterium, 599 neurocysticercosis, 2474, 2476, 2477 neurosyphilis, 1059, 1061–1062, 1074 non-spore-forming, anaerobic, Grampositive rods, 923 Orientia, 1124 parasitology, 2294, 2298 Parastrongylus, 2498 phaeohyphomycoses, 2161, 2163–2164 progressive multifocal leukoencephalopathy (PML), 1407, 1804–1806, 1808, 1811–1812 rabies virus, 1635 Rothia mucilaginosa, 361 scorpion venom, 2520 Spirometra, 2503 Strongyloides stercoralis, 2457 Toxoplasma gondii, 2375 transmissible spongiform encephalopathies (TSEs), 1859–1864 Tropheryma whipplei, 161, 1159 Trypanosoma brucei, 2366–2367 varicella-zoster virus, 1705–1706, 1709 Central nervous system samples parasitology, 2298 rabies virus, 1636–1640 Central nervous system toxicity, of penicillins, 1173 Central-line-associated bloodstream infection (CLABSI), 107 Centrifugation-enhanced rapid cell culture, 1426–1427 Centrifuges, 174 Centrocestus, 2482 Centruroides, 2520 cepA gene, 1346 Cephalexin, 1173, 1198 Cephalosporin(s), 1173–1176 adverse effects, 1175–1176 antimicrobial susceptibility testing, 1255, 1259 concentration in serum, 1198 expanded-spectrum, 1174–1175, 1324 first-generation, 1174 fourth-generation, 1175 mechanism of action, 1173 pharmacology, 1173 second-generation (broad-spectrum), 1174 spectrum of activity, 1174–1175 structure, 1172 third-generation, 1174 Cephalosporin resistance, 1173–1175 Bacillus, 1326 Bacteroides fragilis group, 1346 Clostridium difficile, 1348 Fusobacterium mortiferum, 1348 Haemophilus influenzae, 1321 Neisseria meningitidis, 1324 Streptococcus pneumoniae, 1316 Cephalosporinases, 1223, 1227 Cephalosporium, 2071 Cephalosporium acremonium, 1173 Cephalotheca, 2062, 2069, 2075 Cephalotheca foveolata, 2062, 2069, 2070 Cephamycins, 1174 Cepheid CT/NG Xpert Rapid PCR test, 639 Cepheid Expert HPV assay, 1790 Cepheid GeneXpert MTB/RIF, 1356

SUBJECT INDEX Cepheid RSV, 1506 Cercarial dermatitis, 2484 Cercopithecine herpesvirus 1, see Herpes B virus Cerebellar ataxia, varicella-zoster virus and, 1709 Cerebellitis, varicella-zoster virus and, 1709 Cerebral amyloid angiopathy, 1859 Cerebral atrophy, JC polyomavirus and, 1804 Cerebral infection homothallic ascomycetes, 2075 mucormycosis, 2089 Cerebriform, 1941 Cerebrospinal fluid (CSF) specimen amebae, 2392–2393, 2395 Anaplasma phagocyrophilum, 1139–1140 arboviruses, 1648 Borrelia, 1042, 1047 Brucella, 865 Candida, 1996 Cryptococcus, 1977, 1993, 1997 Ehrlichia chaffeensis, 1139–1140 enterovirus, 1541–1542 Epstein-Barr virus, 1741, 1747 fungi, 1944, 1946–1947, 1950, 1953 Gram stain and plating medium recommendations, 286 Haemophilus and, 670–672 herpes simplex virus (HSV), 1689–1690, 1695–1696, 1696 Histoplasma capsulatum, 1977, 2116, 2122 human herpesvirus 6 (HHV-6), 1754, 1760 human T-cell lymphotropic viruses (HTLVs), 1460 Leptospira, 1031 mumps virus, 1493 parasitology, 2327–2328 Parastrongylus, 2498 polyomavirus, 1806–1807, 1812 prions, 1862 rabies virus, 1636, 1641 specimen collection, transport, and storage guidelines, 273, 294 streptococcal antigen detection in, 388 Streptococcus pneumoniae, 1319 Toxoplasma gondii, 2375, 2380–2381 transmissible spongiform encephalopathies (TSEs), 1864 Treponema, 1062, 1073–1074 Trypanosoma brucei, 2366–2368 varicella-zoster virus, 1707–1709, 1712–1713 viruses, collection methods and processing of specimens, 1413 Cerinosterus cyanescens, 2071 Ceriporia lacerata, 2071, 2075 Cervarix, 1786 Cervical adenopathy, 1771 Cervical cancer human papillomavirus, 1784, 1786, 1788 screening recommendation, 1786 Cervical lymphadenitis Mycobacterium haemophilum, 542 Mycobacterium scrofulaceum, 544 Mycobacterium szulgai, 543 Mycobacterium tuberculosis, 538 Cervical secretion specimens, 277 Cervicitis Chlamydia trachomatis, 1108 etiologies, usual, 290 Mycoplasma, 1092, 1101 Trichomonas vaginalis, 2414 viruses, specimens and methods for detection of, 1408

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 57

Cervista, 1414 Cervista HPV genotyping assay, 1794 Cervista HR-HPV test, 1792–1793 Cestoda (class), 2471, 2473–2475, 2501–2502 Cestodes (tapeworms), 2471–2477 detection, 2323 Diphyllobothrium latum, 2471–2473 Echinococcus granulosus, 2471, 2476–2477 Echinococcus multilocularis, 2471, 2476–2477 Hymenolepis nana, 2471–2472, 2475–2476 less common, 2499–2503 Taenia multiceps, 2471, 2477 Taenia saginata, 2471–2474 Taenia serialis, 2477 Taenia solium, 2471–2472, 2474–2475 taxonomy and classification, 2288, 2291 treatment, 2531, 2533 Cestodiasis, 2476 Cethromycin, 1198 Cetrimide agar, non-USP, 332 Cetrimide agar, USP (Pseudosel agar), 332 Cetylpyridinium chloride-sodium chloride (CPC-NaCl), 320 cfiA gene, 1346–1347, 1383 cfr gene, 1385 cfxA gene, 1346 Chabertiidae (family), 2289 Chaetocladiaceae (family), 2087 Chaetomium, 2062, 2069, 2075, 2161, 2192 Chaetomium atrobrunneum, 2062 Chaetomium globosum, 2062, 2075, 2155, 2160 Chaetomium perlucidum, 2062, 2155 Chaetosartorya, 2030 Chaetothyriales (order), 1938, 2153–2154, 2156–2159, 2162–2163, 2165–2166, 2173–2174 Chagas’ disease, 2357, 2362–2365; see also Trypanosoma cruzi arthropod vector, 2507–2508 commercial kits for immunodetection of serum antibodies, 2296 detection, 2331, 2364–2365 treatment, 2365, 2530, 2545 xenodiagnosis, 2307, 2365 Chagoma, 2362 Chancre Treponema, 1059 Trypanosoma brucei, 2366 Chancroid, 670–671 Chapare virus, 1669, 1672, 1674 Charcoal cefoperazone deoxycholate agar, 1002–1003 Charcoal-based selective medium (CSM), 1002–1003 Check HMPV assay, 1510 Check-MDR assay, 1383 Check-Points, 1383–1384 Cheiracanthium inclusum, 2521 Cheiracanthium mildei, 2521 Chelicerae, 2512, 2520 Chembio Dual Path Platform, 1070, 1072, 1445 Chemical fume protection, 172–173 Chemiluminescence immunoassay (CLA), 100–101 Epstein-Barr virus, 1743–1744 hepatitis C virus, 1607–1608 human immunodeficiency virus, 1444 human T-cell lymphotropic viruses (HTLVs), 1462–1463, 1465

n lvii

Treponema, 1071–1072 Trypanosoma cruzi, 2365 Chemotaxonomy, 262 Chest pain dirofilariasis, 2499 Histoplasma capsulatum, 2114 Paragonimus, 2487 Chickenpox, 1704–1706; see also Varicella-zoster virus clinical features, 1704–1706 complications, 1704–1705 laboratory tests suggested for, 125 vaccine, 1704, 1706, 1709 Chiggers, 2511 Chigoe, 2509–2510 Chikungunya virus, 1644, 1646, 1651–1652 Childbed fever, 385 Chills arenaviruses, 1673 Coccidioides, 2114 filoviruses, 1674 Histoplasma capsulatum, 2114 malaria, 2339 parvovirus B19, 1819 spider envenomation, 2520 Chilomastix, 2408 Chilomastix mesnili, 2321, 2400, 2408–2410, 2416 Chilopoda (class), 2522 Chlamydia, 1106–1117 antimicrobial susceptibilities, 1116, 1184, 1195 detection with fluorescent antibody, 1423–1425 diagnostic tests, 1084 in Acanthamoeba, 2389 rapid cell culture, 1426 Chlamydia abortus, 1107 clinical significance, epidemiology, and transmission, 1109 epidemiology and clinical diseases associated with, 1083 genome, 1108 Chlamydia caviae, 1108–1109 Chlamydia felis, 1108–1109 Chlamydia isolation medium, 1430 Chlamydia muridarum, 1108–1109 Chlamydia pecorum, 1106 Chlamydia pneumoniae, 1091 antimicrobial susceptibilities, 1116, 1180, 1183, 1187, 1190 biosafety considerations, 1113 clinical significance, epidemiology, and transmission, 1108–1109 complement fixation, 1115 diagnostic tests, 1084 enzyme immunoassay (EIA), 1116 epidemiology and clinical diseases associated with, 1083 evaluation, interpretation, and reporting of results, 1117 genome, 1108 identification, 1107, 1114–1115 isolation, 1113–1114 MIF test, 1115–1116 serologic tests, 1084, 1115–1116 specimen collection, transport, and handling, 299, 1111 taxonomy, 1106 Chlamydia psittaci antimicrobial susceptibilities, 1116 biosafety considerations, 1113 biothreat agent, 223 clinical significance, epidemiology, and transmission, 1109

lviii

n

SUBJECT INDEX

Chlamydia psittaci (continued) diagnostic tests, 1084 epidemiology and clinical diseases associated with, 1083 evaluation, interpretation, and reporting of results, 1117 genome, 1108 identification, 1114–1115 isolation, 1113–1114 MIF test, 1115–1116 specimen collection, transport, and handling, 299, 1111 taxonomy, 1106 transmission and disease, 223 typing systems, 1115 Chlamydia suis, 1109 Chlamydia trachomatis, 1091, 2505 antimicrobial susceptibilities, 1116, 1180, 1183, 1186–1187 biosafety considerations, 1113 clinical significance, epidemiology, and transmission, 1108 complement fixation, 1115 DFA and IFA reagents for the detection of, 1425 diagnostic tests, 1084 enzyme immunoassay (EIA), 1116 epidemiology and clinical diseases associated with, 1083 evaluation, interpretation, and reporting of results, 1116–1117 genome, 1108 identification, 1107, 1114–1115 immunofluorescence detection of, 1426 isolation, 1113–1114 MIF test, 1115–1116 serologic tests, 1084, 1115–1116 specimen collection, transport, and handling, 295–296, 299, 1110–1111 taxonomy, 1106 typing systems, 1115 Chlamydiaceae (family), 1106–1117 antigen detection assays, 1112–1113 direct fluorescent antibody tests (DFAs), 1112–1113 enzyme immunoassays, 1113 point-of-care tests, 1113 antimicrobial susceptibilities, 1116 clinical significance, epidemiology, and transmission, 1083, 1108–1109 Chlamydia abortus, 1109 Chlamydia pneumoniae, 1108–1109 Chlamydia psittaci, 1109 Chlamydia trachomatis, 1108 environmental chlamydiae, 1109 collection, transport, and storage of specimens, 1109–1111 Chlamydia pneumoniae, 1111 Chlamydia psittaci, 1111 Chlamydia trachomatis, 1110–1111 general comments, 1109–1110 direct examination, 1111–1113 evaluation, interpretation, and reporting of results, 1116–1117 identification, 1114–1115 isolation, 1113–1114 biosafety considerations, 1113 procedures, 1113–1114 specimen processing, 1113 nucleic acid amplification tests (NAATs), 1111–1112 Chlamydia pneumoniae, 1112 Chlamydia psittaci, 1112

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 58

Chlamydia trachomatis, 1111–1112 nucleic acid hybridization tests, 1112 serologic tests, 1084, 1115–1116 complement fixation, 1115 enzyme immunoassay (EIA), 1116 MIF test, 1115–1116 specimen processing, 1113 blood, 1113 bubo pus, 1113 fecal samples, 1113 ocular genital tract specimens, 1113 sputum, throat washings, respiratory tract secretions, 1113 tissue samples, 1113 taxonomy, 1106 typing systems, 1115 Chlamydiales (order), 1106 Chlamydia-like endosymbiont, 1108–1109 Chlamydophila, see Chlamydia Chlamydophila pneumophila, in Acanthamoeba, 2389 Chlamydospores, 1939, 1941, 2060–2061 Chlamylege, 892 Chloramphenicol, 1193–1194 adverse effects, 1193–1194 antimicrobial susceptibility testing, 1256, 1261 concentration in serum, 1198 fungi inhibition, 1949, 1951–1952 mechanism of action, 1193 pharmacology, 1193 spectrum of activity, 1193 Chloramphenicol acetyltransferases (CATs), 1217, 1228 Chloramphenicol resistance, 1228–1229 acetyltransferases, 1228 common associations of resistance mechanisms, 1215 decreased accumulation of chloramphenicol, 1228–1229 Chlorhexidine, 185 Acanthamoeba, 2394 for decolonization of MRSA, 188–189 presurgical skin disinfection, 188 skin disinfection, 17 Chlorine, 193 Chlorine-releasing compounds, 193 Chloroquine, 2349, 2536–2537, 2564–2565 adverse effects, 2537 mechanism of action, 2536 pharmacokinetics, 2537 spectrum of activity, 2537 Chloroquine resistance, 2550–2553, 2564–2565 Chocolate agar, 332 Chocolate tellurite agar (tellurite blood agar), 332 Cholangiopathy, microsporidia and, 2210, 2213 Cholangitis Ascaris lumbricoides, 2451 Bacillus clausii, 442 liver trematodes, 2489 Stenotrophomonas maltophilia, 794 Cholecystitis Aeromonas, 754 Ascaris lumbricoides, 2451 Campylobacter, 1000 Cellulomonas, 479 cephalosporins, 1175 Cyclospora cayetanensis, 2429 Dolosigranulum pigrum, 424 Lactococcus, 424 liver trematodes, 2489

microsporidia, 2210 non-spore-forming, anaerobic, Gram-positive rods, 923 Vibrio metschnikovii, 766 Cholelithiasis, liver trematodes, 2489 Cholera, 125, 762–763 Cholera and Other Vibrio Illness Surveillance (COVIS) system, 769 Cholera medium (thiosulfate-citrate-bile salts-sucrose [TCBS] agar), 332 Cholera medium base with tellurite and blood, 332 Chopped meat broth, 332 Chordopoxvirinae (subfamily), 1398, 1828, 1834 Chorioamnionitis, Facklamia, 425 Chorioretinal lesions, Onchocerca volvulus, 2466 Chorioretinitis, Toxoplasma gondii, 2381 Christensen agar, 332; see also Urea agar Christensenella minuta, 969 Christensenellaceae (family), 969 Christensen’s urea agar, 1960 Chromadorea (class), 2467 CHROMagar, 1960, 1986 CHROMagar Acinetobacter, 332 CHROMagar Candida, 1952, 1991, 1999 CHROMagar CTX (RambaCHROM CTX), 332 CHROMagar E. coli (RambaCHROM E. coli), 332 CHROMagar ECC (RambaCHROM ECC), 333 CHROMagar ESBL (RambaCHROM ESBL), 333 CHROMagar KPC (RambaCHROM KPC), 333 CHROMagar Listeria (RambaCHROM Listeria), 333 CHROMagar MRSA (RambaCHROM MRSA), 333 CHROMagar MRSA II, 1293 CHROMagar O157 (RambaCHROM O157), 333 CHROMagar Orientation (RambaCHROM Orientation), 333 CHROMagar Pseudomonas (RambaCHROM Pseudomonas), 333 CHROMagar Salmonella (RambaCHROM Salmonella), 333 CHROMagar Salmonella Plus (RambaCHROM Salmonella Plus), 333 CHROMagar Staph Aureus (RambaCHROM Staph Aureus), 333 CHROMagar STEC (RambaCHROM STEC), 333 CHROMagar StrepB (RambaCHROM StrepB), 333 CHROMagar Vibrio (RambaCHROM Vibrio), 333 CHROMagar VRE (RambaCHROM VRE), 333 CHROMagar Y. enterocolitica (RambaCHROM Y. enterocolitica), 333 Chromatic aberration, 5 Chromatographic immunoassay (CIA) influenza viruses, 1472–1475 respiratory syncytial virus, 1502, 1504 chromID C. difficile, 334 chromID Candida, 1999 chromID Candida agar (CAN2) (bioMérieux), 1960

SUBJECT INDEX chromID CARBA agar, 334 chromID CPS, 334 chromID ESBL, 334 chromID MRSA, 334, 1293 chromID P. aeruginosa, 334 chromID S. aureus, 334 chromID Salmonella, 334 chromID Strepto B, 334 chromID Vibrio, 334 chromID VRE, 334 Chromista (kingdom), 1939, 2286–2287, 2406 Chromobacterium, 652 antimicrobial susceptibilities, 662 clinical significance, 654 epidemiology and transmission, 653–654 identification, 658–659 isolation procedures, 656 serotyping, 661 taxonomy and description of, 653 Chromobacterium haemolyticum, 653 Chromobacterium violaceum, 653, 656, 661–662 Chromoblastomycosis, 1975, 2153, 2156 antifungal susceptibilities, 2167 clinical significance, 2163–2164 epidemiology and transmission, 2161–2162 microscopy, 2164 Chromoblastosis, specimens for, 1947 Chromogenic agars for oxacillin resistance detection in Staphylococcus, 1293 for yeast identification, 1999 Chromogenic enzyme substrate tests, Neisseria, 641 Chronic aspiration syndrome, Mycobacterium, 599 Chronic fatigue syndrome, Chlamydia pneumoniae, 1109 Chronic granulomatous disease Burkholderia, 793 Pandoraea, 795 Chronic obstructive pulmonary disease (COPD) Pandoraea, 795 respiratory syncytial virus (RSV), 1500 rhinoviruses, 1553 Chryseobacterium, 616 antimicrobial susceptibilities, 829 identification, 827–828 odor of, 823 taxonomy, 813 Chryseobacterium anthropi, 624–625, 827–829 Chryseobacterium caeni, 614 Chryseobacterium gleum, 624–625, 826–828 Chryseobacterium hominis, 624–625, 827–828 Chryseobacterium indologenes, 624–625, 826–828 Chryseobacterium treverense, 624–625, 826, 828 Chrysomia, 2330 Chrysops, 2507 Chrysosporium, 2063, 2069, 2073, 2076, 2109, 2118, 2139 Chrysosporium guarroi, 2076 Chrysosporium ophiodiicola, 2063, 2076 Chrysosporium zonatum, 2063, 2073, 2076 Ciclopirox, for dermatophytes, 2145 Cidofovir adenoviruses, 1777 antiviral susceptibility testing, 1916 cytomegalovirus, 1720 enteric adenoviruses, 1622

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 59

herpes simplex virus (HSV), 1689, 1695 herpesviruses, 1883, 1885 human herpesvirus 6 (HHV-6), 1760 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1763 orthopoxviruses, 1832 polyomavirus, 1811 polyomavirus-associated nephropathy (PVAN), 1811 progressive multifocal leukoencephalopathy (PML), 1811 varicella-zoster virus, 1712 Cidofovir resistance, 1917 cytomegalovirus, 1730, 1895–1896 herpes simplex virus (HSV), 1894–1895 varicella-zoster virus, 1895 Ciliates, 2400, 2416–2417 Ciliophora (phylum), 2287, 2416 Cimex hemipterus, 2508 Cimex lectularius, 2508 Cinchona alkaloids, 2537–2538 adverse effects, 2538 mechanism of action, 2537 pharmacokinetics, 2537–2538 spectrum of activity, 2538 CINtec p16, 1789 CINtec Plus, 1789 Ciprofloxacin, 1178–1180, 1198 antimicrobial susceptibility testing, 1256, 1260 Cyclospora cayetanensis, 2431 Cystoisospora belli, 2431 Ciprofloxacin resistance, 1214 Bacillus anthracis, 1325 Francisella tularensis, 1325 Neisseria gonorrhoeae, 1323 Yersinia pestis, 1325 Circulating cathodic antigen, 2486 Circulating recombinant form (CRF), HIV, 1436, 1446 Circulatory system, trematodes of, 2479–2486 Cirrhosis, liver trematodes and, 2489 Citricoccus, 354, 361 Citrinin, 2189, 2191 Citrobacter antimicrobial susceptibilities, 727–729, 1177–1178, 1180, 1196 epidemiology, transmission, and clinical significance, 720 identification, 725, 727 identification of species, 32 separation of members of genus, 723 Citrobacter amalonaticus, 716, 723, 725 Citrobacter braakii, 716, 720, 723, 725 Citrobacter farmeri, 716, 723 Citrobacter freundii, 716, 720, 723, 725, 728–729 antibiotic resistance, 1224, 1226–1227 β-lactamases, 1299 Citrobacter gillenii, 716, 720, 723, 725 Citrobacter koseri, 716, 720, 723, 725 Citrobacter murliniae, 716, 720, 723 Citrobacter rodentium, 716, 720, 723 Citrobacter sedlakii, 716, 723, 725 Citrobacter werkmanii, 32, 716, 723, 725 Citrobacter youngae, 716, 723, 725 CJD, see Creutzfeldt-Jakob disease Cladophialophora, 1939, 1940, 2153–2154, 2156, 2165–2166, 2168 Cladophialophora arxii, 2154, 2163 Cladophialophora bantiana, 2154, 2156–2157, 2161–2163, 2165, 2167, 2173, 2268–2269

n lix

Cladophialophora boppii, 2154, 2162–2163 Cladophialophora carrionii, 1967, 2154, 2156, 2161, 2163, 2167 Cladophialophora devriesii, 2154, 2163 Cladophialophora emmonsii, 2154, 2162 Cladophialophora modesta, 2154, 2163 Cladophialophora mycetomatis, 2154 Cladophialophora samoënsis, 2154, 2163 Cladophialophora saturnica, 2154, 2162 Cladophialophora yegresii, 2161 Cladosporium, 2154–2155, 2162, 2192 Cladosporium cladosporioides, 1991, 2154–2155, 2158, 2162 Cladosporium oxysporum, 2154–2155, 2162 Cladosporium werneckii, 2147 ClariRes assay, 1020, 1023 Clarithromycin, 1182–1183, 1198 antimicrobial susceptibility testing, 1255, 1260 Balamuthia mandrillaris, 2395 for Mycobacterium, 1361 Toxoplasma gondii, 2382 Clarithromycin resistance, 1231, 1329 CLART HPV2, 1791 CLART PneumoVir, 1576 Classification of bacteria, 255–265 criteria for species delineation, 256 genomic threshold for species definition, 257–258 major groups of bacteria, 258–259 methods, 259–263 amplified fragment length polymorphism (AFLP), 261 cell wall composition, 262 cellular fatty acid analysis, 262 chemical methods, 262 DNA-DNA hybridization studies, 259– 260 FTIR spectroscopy, 263 G+C content, 261 mass spectrometry, 262–263 PCR-based, 261 phenotypic methods, 261–262 rRNA studies, 260–261 sequence analysis of protein-encoding genes, 261 whole-genome sequence-based methods, 260 multilocus sequence analysis, 257 polyphasic species concept, 256–257 uncultured bacteria, 259 Classification of viruses, 1393–1402 Clavate, 1941 Claviceps, 2190 Claviceps purpurea, 2190 Clavines, 2190 Clavispora, 1938 Clavispora lusitaniae, 1985, 1993, 2000 Clavulanic acid, 1177 with amoxicillin, 1177, 1198 ESBL inhibition by, 1299 with ticarcillin, 1177, 1199 Cleaning decontamination, 189–190 definition, 190 surfaces and floors, 196–197 Clearview Exact Influenza A & B, 1474 Clearview RSV, 1504 Cleavase-Invader technology, 56–57, 58 Cleistothecia, 1941, 2033, 2041–2042, 2180 CLIA (Clinical Laboratory Improvement Amendments of 1988), 31, 217, 1281 Climate change, mycotoxins and, 2191

lx

n

SUBJECT INDEX

Clindamycin adverse effects, 1185 anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1256, 1261 Bacteroides fragilis group susceptibility percentages, 1350 concentration in serum, 1198 for Mycobacterium avium complex, 1369– 1370 mechanism of action, 1185 pharmacology, 1185 spectrum of activity, 1185 Toxoplasma gondii, 2382 Clindamycin resistance, 1231 Bacteroides fragilis group, 1347 Clostridium difficile, 1348 Clostridium perfringens, 1348 detection in Staphylococcus, 1290, 1295– 1297 D-zone test, 1267, 1296–1297 quality control and quality assessment, 1296 reporting of results, 1296 single-well broth dilution method, 1267, 1296 detection in Streptococcus D-zone test, 1267, 1297–1298 quality control and quality assessment, 1298 reporting of results, 1298 single-well broth dilution method, 1267, 1298 Gram-positive cocci, 1348 inducible, 1279, 1290, 1295–1298 Porphyromonas, 1347 Streptococcus, 1320 Streptococcus pneumoniae, 1297–1298, 1316 Clinical and Laboratory Standards Institute (CLSI), 33, 108, 183 antifungal susceptibility testing, 2255–2273 antimicrobial susceptibility testing anaerobic bacteria, 1342–1345, 1347– 1352 Aspergillus, 2044–2045 automated systems, 1274, 1277–1280 breakpoints, 1248–1249 Campylobacter, 1007 clinical and bacteriological response rates, 1248 confirmatory and supplementary test use, 1250 diffusion testing, 1264–1267 dilution testing, 1254–1264 Escherichia coli, 696 fastidious bacteria, 1319–1332 Mycobacterium, 1356, 1362–1363 Pasteurella, 661 Pseudomonas, 783 quality control, 1264, 1267 reference methods, 1253 selection of antibacterial agents for testing, 1247, 1254 selection of testing method, 1247 Staphylococcus, 369–370 “susceptible dose dependent” interpretive category, 1246 phenotypic methods for detecting antibacterial resistance, 1286–1303 website, 1249 Clinical laboratory design, biosafety and, 171–172

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 60

Clinical Laboratory Improvement Amendments of 1988 (CLIA), 31, 217, 1281 Clinical microbiology laboratory, 44–53 automation automated specimen processing, 48–49 digital imaging, 51 evaluation and selection criteria for system, 51–52 future perspectives, 52 historical perspective, 47–48 limitations of systems, 51 molecular automation, 48 organism identification, 48 susceptibility testing, 48 total laboratory automation (TLA), 49– 51 geography, 44–45 infectious disease testing, 45 location of testing, 45 preanalytical microbiology, 45 quality control, 45 staff training level, 46 staffing models, 45–46 workflow, 46–47 batch versus immediate testing, 46–47 process improvement, 47 Clinostomidae (family), 2290 CLIP sequencing, 69 Clofazimine activity, 1360 adverse effects, 1360 for Mycobacterium infection, 1360 Clonal relatedness of isolates, 148–149 Clone, definition, 132 Clonorchiasis, 2489–2490, 2533 Clonorchis, 2320, 2328, 2330 Clonorchis sinensis, 2449, 2481, 2484, 2488–2490, 2531 Clostridiaceae (family), 922 Clostridial agar, 334 Clostridiales (order), 922 Clostridium, 940–959 antibiotic resistance, 1234, 1348 antimicrobial susceptibilities, 957–958, 1172, 1175, 1177, 1180, 1184, 1187–1190, 1194, 1348, 1351 antimicrobial susceptibility testing, 1345 characteristics of species, 942–943 clinical significance, 941–948 bacteremia, 941–942 botulism, 946–947 C. difficile infection, 944 enteric infections, 942–944 enteritis necroticans, necrotizing enteritis, necrotizing enterocolitis (NEC), 943–944 food poisoning, 942–943 gynecologic infections, 945 histotoxic skin and soft tissue infections, 944–946 myonecrosis, 945 tetanus, 947–948 collection, transport, and storage of clinical specimens, 948–952 suspected C. botulinum or C. tetani infection or intoxication, 951– 952 suspected C. difficile infection, 949–951 suspected C. perfringens foodborne illness, 948 suspected enteritis necroticans (C. perfringens type C), 948–949

suspected gas gangrene or necrotizing fasciitis, 948 suspected neutropenic enterocolitis involving C. septicum, 951 culture of, 922 description of genus, 940–941 epidemiology and transmission, 941 evaluation, interpretation, and reporting of results, 957, 959 genome sequences, 941 identification, 953–956 carbohydrate fermentation, 954–955 characteristics of commonly encountered clostridia, 955–956 definitive, 954–955 esculin hydrolysis, 954 gelatin hydrolysis, 954 lecithinase, 953–954 lipase, 953–954 PRAS media, 954 preliminary, 953–954 special potency disks, 953 spore test, 953–954 isolation procedures, 952–953 C. difficile, 953 on plated media, 952 spore selection techniques, 952–953 serologic tests, 957 taxonomy, 940 typing systems, 956–957 vanB gene, 1381 Clostridium argentinense, 942–943, 946 Clostridium baratii, 942–943, 946 Clostridium bifermentans, 941–943, 954–955 Clostridium bolteae, 940, 942–943, 948, 955, 958 Clostridium botulinum antimicrobial susceptibilities, 958 biothreat agent, 221 characteristics, 942–943 clinical significance, 946–947 collection, transport, and storage of clinical specimens, 303, 951–952 genome sequence, 941 identification, 955–956 spore selection techniques, 952 toxins, 946–947, 952 transmission and disease, 221 typing, 957 Clostridium butyricum, 940, 942–943, 946, 955, 957–958 Clostridium cadaveris, 942–943, 955, 958, 1348 Clostridium carnis, 940, 942–943 Clostridium celerecrescens, 948 Clostridium chauvoei, 955 Clostridium clostridioforme, 940, 942–943, 948, 953, 955–958, 1348 Clostridium coccoides, 940 Clostridium cocleatum, 940 Clostridium difficile antimicrobial susceptibilities, 957–958, 1175, 1183, 1185, 1188–1190, 1194–1195, 1348, 1351 antimicrobial susceptibility testing, 1343 cell cytotoxicity assays, 1429 characteristics, 942–943, 955 clinical significance, 905, 941, 944 pathogenicity locus, 944 prevalence of C. difficile infections, 944 risk factors and course of C. difficile infections, 944 colitis, 944, 1180, 1185, 1188, 1194 commercial sources of chromogenic agar media for, 326

SUBJECT INDEX disinfection, 192–193, 196, 198 epidemiology and transmission, 941 evaluation, interpretation, and reporting of results, 959 genome sequence, 941 health care-associated infections, 115–116 isolation, 953 reference strains, 1343–1344 specimen collection, transport, and storage guidelines, 275, 301–302, 949–951 toxigenic culture, 906 toxins, 944, 948 typing systems, 956–957 Clostridium disporicum, 958 Clostridium fallax, 948, 955 Clostridium ghonii, 922 Clostridium glycolicum, 942–943, 955, 958 Clostridium hastiforme, 942–943 Clostridium hathewayi, 940, 942–943, 948, 955, 958 Clostridium histolyticum, 940, 942–945 Clostridium indolis, 942–943 Clostridium innocuum, 940, 942–943, 948, 953, 956–958, 1183, 1348 Clostridium intestinale, 948 Clostridium leptum, 922, 940 Clostridium limosum, 942–943 Clostridium neonatale, 940, 944 Clostridium novyi, 942–945, 954, 956 Clostridium orbiscindens, 922; see also Flavonifractor plautii Clostridium paraputrificum, 942–943, 958 Clostridium perfringens, 976 antibiotic resistance, 1228, 1348 antimicrobial susceptibilities, 957–958, 1183–1185, 1189–1190, 1348, 1351, 2535 antimicrobial susceptibility testing, 1343, 1345, 1352 biothreat agent, 223 characteristics, 942–943, 956 clinical significance antibiotic-associated diarrhea, 943–944 bacteremia, 941–942 enteritis necroticans and necrotizing enterocolitis, 943–944 food poisoning, 942–943, 948 collection, transport, and storage of clinical specimens, 948 colony morphology, 952 culturing, 906 description, 940–941 enterotoxin, 942–943 epidemiology and transmission, 941 evaluation, interpretation, and reporting of results, 957, 959 extracellular toxins, 945–946 genome sequence, 941 identification, 953–954 isolation procedures, 952 spore selection techniques, 952 thioglycolate bile broth, 345 transmission and disease, 223 type A, 942–944 type C, 943, 948–949 typing, 957 Clostridium putrificum, 942–943 Clostridium ramosum, 940, 942–943, 948, 953, 956–958, 1183, 1348 Clostridium septicum, 941–946, 951–958, 1352 Clostridium sordellii, 922, 941–946, 954, 958– 959, 1352

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 61

Clostridium sphenoides, 942–943 Clostridium spiroforme, 940 Clostridium sporogenes, 942–943, 954, 956–959 Clostridium subterminale, 942–943, 957–958, 1348 Clostridium symbiosum, 942–943, 948, 956 Clostridium tertium, 940–943, 948, 956–958 Clostridium tetani antimicrobial susceptibilities, 958 characteristics, 942–943, 956 clinical significance, 947–948 collection, transport, and storage of clinical specimens, 951–952 description, 940 genome sequence, 941 identification, 954 toxins, 947–948, 952 CLOtest, 1019 Clotrimazole, 2394 Cloxacillin, 1171–1172, 1198, 1299–1300 CLSI, see Clinical and Laboratory Standards Institute Cluster, definition, 132 Cluster detection, molecular surveillance and, 145–146 Clustered regularly interspaced short palindromic repeats (CRISPRs) analysis, 140–141 CMV, see Cytomegalovirus CMV Brite, 1724 CMV Brite Turbo, 1724 CMV pp65 antigenemia assay, 1413 CMV TurboTreat medium, 1430 CMX001, 1777, 1832 CNS, see Central nervous system disease/ infection Coagulase production, by Staphylococcus, 362–363 Coagulase test, 317, 352 Coagulase-mannitol agar, 334 Coagulopathy cephalosporins, 1175 herpes simplex virus (HSV), 1689 Cobas 4800 CT/NG test, 639, 1110–1111 Cobas 4800 system, 74 Cobas Amplicor CMV Monitor, 1726 Cobas Amplicor HCV test, 1411 Cobas Amplicor HIV-1 test, 1411 Cobas Amplicor Monitor, 1610 Cobas Amplicor v2.0, 1603 Cobas AmpliPrep system, 74 Cobas AmpliPrep/Cobas Amplicor v2.0, 1603 Cobas AmpliPrep/Cobas TaqMan, 74, 1604– 1605, 1610 Cobas AmpliPrep/Cobas TaqMan CMV test, 1411, 1725–1726 Cobas AmpliPrep/Cobas TaqMan HBV test, 1411 Cobas AmpliPrep/Cobas TaqMan HCV test, 1411 Cobas AmpliPrep/Cobas TaqMan HIV-1 test, 1411, 1442–1443 Cobas HBsAg, 1847 Cobas HPV, 1414 Cobas HPV test, 1786–1787, 1793 Cobas TaqMan, 74, 1610–1611 Cobas TaqMan HBV, 1849 Cobicistat, 1872, 1877–1878 Cocci Gram-negative anaerobic cocci (GNAC), 909–916 Gram-positive anaerobic cocci (GPAC), 909–916

n lxi

Gram-positive cocci antibacterial resistance patterns, 1348– 1349, 1352 catalase-negative, 422–431 catalase-positive, 354–372 identification of aerobic, 350–352 Coccidea (class), 2287, 2373, 2425, 2435 Coccidia, 2425–2431 antigen detection, 2430 clinical significance, 2428–2429 collection, transport, and storage of specimens, 2429 culture, 2430 description of agents, 2425–2427 detection, 2325 direct examination, 2429–2430 epidemiology, transmission, and prevention, 2427–2428 evaluation, interpretation, and reporting of results, 2431 life cycles, 2425–2427 microscopy, 2426–2427, 2429–2430 nucleic acid detection, 2430 serologic tests, 2430 taxonomy, 2425 treatment, 2430–2431 Coccidiasina (subclass), 2373 Coccidioides, 895, 1939, 1994, 2109–2123 antibody detection, 1971 antifungal susceptibilities, 2121–2122, 2224 antigen detection, 1977, 2116 biosafety, 2117 biosecurity, 2117 clinical significance, 2114 description of agents, 2111–2112 direct examination, 2115–2117 endophthalmitis, 1949 epidemiology and transmission, 2113 evaluation, interpretation, and reporting of results, 2123 identification, 2118–2119 isolation, 2117–2118 laboratory-acquired infections, 177 microscopy, 1969, 2112, 2116 nucleic acid detection, 2117 serologic tests, 2121 specimen collection, transport, and processing, 1948–1949, 1951, 2115 taxonomy, 2110 typing systems, 2120 Coccidioides immitis, 1424, 1937, 1966, 1976, 2109–2110, 2112, 2120 Coccidioides posadasii, 1937, 1966, 2109–2110, 2120 Coccidioidomycosis, 2109–2123 agent, 2110 antigen detection, 2116 clinical significance, 2114 collection, transport, and storage of specimens, 1947, 2115 epidemiology and transmission, 2113 evaluation, interpretation, and reporting of results, 2123 nucleic acid detection, 2117 serologic tests, 2121 typing, 2119 Cochlear implant-related infection, Nocardia farcinica, 517 Cochliobolus australiensis, 2155 Cochliobolus geniculatus, 2155 Cochliobolus hawaiiensis, 2155 Cochliobolus lunatus, 2155 Cochliobolus spicifera, 2155

lxii

n

SUBJECT INDEX

Cochliomyia, 2330, 2519 Cockroaches, 2513, 2515, 2522 Coelomycetes, 1939, 1941, 2065, 2075–2076, 2077 Coenurosis, 2471, 2477 Coenurus, 2329, 2332, 2477 Cognitive disturbances, mucormycosis and, 2089 Cohnella hongkongensis, 443 Cokeromyces, 2088, 2093–2094 Cokeromyces recurvatus, 2088, 2093–2094, 2098 ColdTrack vials, 164 Coleophoma empetri, 2228 Coleoptera (order), 2521–2522 Colistimethate sodium, 1198 Colistin, 616, 1192 Colistin resistance, 1232 Colistin-oxolinic acid-blood agar, 334 Colitis BK polyomavirus, 1805 Entamoeba histolytica, 2403 Klebsiella oxytoca, 718 Collarette, 1941 Collection, transport, and storage of specimens, see Specimen collection, transport, and processing; specific organisms Collection swabs, for viral culture, 1429 College of American Pathologists, 219 Colletotrichum, 2065, 2069, 2075, 2076 Colletotrichum coccodes, 2065 Colletotrichum crassipes, 2065 Colletotrichum dematium, 2065 Colletotrichum gloeosporioides, 2065 Collierville virus, 1672 Collinsella, 920–921 Collinsella aerofaciens, 930 Collinsella intestinalis, 930 Collinsella stercoris, 930 Colloidal carbon wet mounts, 1957 Colonic microbiome, 230–231 Colorado tick fever virus, 1645, 1648, 25076 Colorimetric antifungal susceptibility testing molds, 2271–2272 yeasts, 2264–2265 Colorimetric microtiter plate (CMP) systems, 67 ColorPAC Giardia assay, 2411 ColorPAC Giardia/Cryptosporidium rapid assay, 2441 Coltivirus (genus), 1399, 1646, 2507 Columbia blood agar, 334 Columbia CNA agar, 334 Columella, 1941 Coma arboviruses, 1647 Rickettsia, 1124 Comamonadaceae (family), 792 Comamonas, 615 characteristics of, 802 clinical significance, 795 description of genus, 792 epidemiology and transmission, 793 identification, 802 taxonomy, 792 Comamonas aquatica, 632–633, 792 Comamonas kerstersii, 632–633, 792 Comamonas terrigena, 630–631, 792, 802 Comamonas testosteroni, 630–631, 795, 802, 804 Combined antiretroviral therapy (cART), 1869 Common cold

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 62

human metapneumovirus, 1509 rhinoviruses, 1553 Community-acquired respiratory distress syndrome (CARDS) toxin, 1088– 1089, 1094 Comparative subtyping, definition, 132 Competitive EIAs, 98 Competitive PCR (cPCR), 73 Complement fixation, 97 Blastomyces, 1969 Blastomyces dermatitidis, 2120 Chlamydiaceae, 1115 Coccidioides, 1971, 2121, 2123 Coxiella burnetii, 1155 Entamoeba histolytica, 2405 fungi, 1969, 1971 Histoplasma capsulatum, 1971, 2120, 2122 influenza viruses, 1481 mumps virus, 1495 Mycoplasma, 1097 Paracoccidioides brasiliensis, 2121, 2123 parainfluenza virus, 1492 respiratory syncytial virus, 1505 Trypanosoma cruzi, 2365 Complete metamorphosis, 2505, 2518–2519 Completoriaceae (family), 2087 Compound microscope, 7–9 Conception product specimen collection, transport, and storage guidelines, 277 Condenser dark-field microscopy, 9 substage, 8 Congenital infection cytomegalovirus, 1719 human herpesvirus 6 (HHV-6), 1755 parvovirus B19, 1819–1820 Toxoplasma gondii, 2373, 2375, 2379–2381 Trypanosoma cruzi, 2363 Congenital rubella syndrome (CRS), 1526–1530 Congenital varicella syndrome, 1412, 1704–1706 Congo red acid morpholinepropanesulfonic acid pigmentation agar, 334 Congo red-brain heart infusion-agarose medium, 334–335 Conidia, 1936, 1938–1941, 2033, 2040–2042, 2058, 2060–2061, 2175 Conidiobolomycosis, 2087, 2099 Conidiobolus, 1937, 1969, 2087, 2099, 2101–2103 Conidiobolus coronatus, 2087, 2102 Conidiobolus incongruus, 2087, 2102 Conidiobolus lamprauges, 2087, 2102–2103 Conidiogenesis, 1939–1940, 1941, 2058 annellidic, 1940 blastic, 1939–1940 phialidic, 1940 thallic, 1939 Conidiogenous cell, 1941 Conidioma, 1941, 2175 Conidiophores, 1941, 2031, 2033, 2040–2042, 2175 Coniochaeta, 2063, 2069, 2071, 2076 Coniochaeta hoffmannii, 2063, 2071, 2076 Coniochaeta mutabilis, 2063, 2071, 2076 Conjugation, 1217 Conjunctiva specimens, 274 Conjunctivitis adenoviruses, 1772 arenaviruses, 1673 Chlamydia trachomatis, 1108 enteric adenoviruses, 1618

etiologies, usual, 290 Haemophilus aegyptius, 670 Haemophilus influenzae, 669 herpes simplex virus (HSV), 1689 Moraxella, 814 Mycobacterium scrofulaceum, 544 Neisseria gonorrhoeae, 636–637 Neisseria meningitidis, 637 Onchocerca volvulus, 2466 Pythium insidiosum, 2201 rubella, 1526 Trypanosoma cruzi, 2362 urticating caterpillar hairs, 2519 urticating spider hairs, 2521 viruses, specimens and methods for detection of, 1407 Conosa (subphylum), 2409 Constipation Anisakis, 2493 Trypanosoma cruzi, 2363 Contagious ecthyma, 1830 Contagious pustular dermatitis, 1830 Contagious pustular stomatitis, 1830 Contaminated products, outbreak investigations on, 126–128 Contamination control, in molecular methods, 78–79 Continuous-monitoring blood culture systems, 21 Contracaecum, 2493 Contrast, 5 Controls, 80 Cooked meat medium, 335 Copan universal transport medium, 1410 Copan Viral Transystem, 1410 COPD, see Chronic obstructive pulmonary disease Copepods, 2507, 2513, 2522 Copiparvovirus (genus), 1818 Copper ions, for disinfection, 195 Coprinus, 2071, 2077 Coprinus cinereus, 2062, 2075 Coprobacter, 967 Coprobacter fastidiosus, 967 Coprococcus eutactus, 922 CoproStrip Giardia/Cryptosporidium, 2441 Cordylobia, 2330 Cordylobia anthropophaga, 2517 Coriobacteriaceae (family), 920 Corn meal agar, 1952 Corneal implant infection, Propionibacterium acnes, 905 Corneal scrapings fungi, 1945, 1949–1950 microsporidia, 2215 specimen collection, transport, and storage guidelines, 275 Corneal ulcers/lesions microsporidia, 2210, 2213 Mycobacterium, 599–600 phaeohyphomycoses, 2162 Pythium insidiosum, 2201 Cornmeal agar with 1% dextrose, 1960 Cornmeal agar with Tween 80, 1960–1961 Coronary artery disease, Helicobacter and, 1018 Coronaviridae (family), 1565, 1618 taxonomic classification, 1398, 1400 virion morphology, 1401 Coronavirinae (subfamily), 1398, 1565 Coronaviruses, 1565–1578 animal, 1565, 1569 antigen detection, 1570 biosafety, 1570, 1577

SUBJECT INDEX clinical significance, 1569 collection, transport, and storage of specimens, 1407–1408, 1569–1570 cytopathic effect (CPE), 1577 description of agent, 1565–1567 detection and identification methods, 1433 direct detection, 1570–1577 discovery, 1566–1567 electron microscopy, 1565, 1570 epidemiology and transmission, 1567–1569 evaluation, interpretation, and reporting of results, 1577–1578 isolation procedures, 1577 microscopy, 1570 natural reservoirs, 1565 nucleic acid detection, 1570–1577 commercial NAATs, 1571, 1574–1576 in-house NAATs, 1571 MERS-CoV, 1577 pan-CoV assays, 1570 SARS-CoV, 1571 species-specific assays, 1571 origin, 1565–1566 phylogenetic relationships, 1566 serologic tests, 1577 structure, 1565–1566 taxonomy, 1565–1566 treatment, 1569 Corynebacterium antimicrobial susceptibilities, 496–497, 1182, 1184, 1188–1190, 1195, 1197 antimicrobial susceptibility testing, 1254, 1317, 1327–1328 blood culture contaminant, 18 chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 479–480 collection, transport, and storage of specimens, 480 description of genus, 474–475 direct examination, 480 epidemiology and transmission, 479 evaluation, interpretation, and reporting of results, 497 identification, 35, 438, 482–494 in skin microbiome, 232 isolation procedures, 480–481 morphologic characteristics, 507 serum tellurite agar for, 344 taxonomy, 474–475, 504–505 typing systems, 496 Corynebacterium accolens, 482, 485, 493 Corynebacterium afermentans, 484, 491, 497 Corynebacterium afermentans subsp. afermentans, 482, 485, 494 Corynebacterium afermentans subsp. lipophilum, 482, 485 Corynebacterium ammoniagenes sensu stricto, 492 Corynebacterium amycolatum, 474, 476, 478–479, 482, 485–486, 489, 492– 494, 497, 1327 Corynebacterium appendicis, 474, 482, 485 Corynebacterium aquaticum, see Leifsonia aquatica Corynebacterium aquatimens, 474, 482, 485 Corynebacterium argentoratense, 482, 485, 487 Corynebacterium atypicum, 474, 482, 487 Corynebacterium aurimucosum, 474, 479, 482, 484, 486–487, 491–492, 494, 497

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 63

Corynebacterium auris, 478, 482, 485, 487, 492, 494 Corynebacterium bovis, 482, 487 Corynebacterium canis, 475, 482, 487 Corynebacterium CDC group F-1, 479, 482, 488, 493 Corynebacterium confusum, 474, 482, 487 Corynebacterium coyleae, 476, 482, 484–485, 487, 491 Corynebacterium diphtheriae, 474, 476, 493 antimicrobial susceptibilities, 1184 antimicrobial susceptibility testing, 1327–1328 CAMP reaction, 481 clinical significance, 479–480 colony morphology, 486 direct examination, 480 epidemiology and transmission, 478 evaluation, interpretation, and reporting of results, 497 Gram stain morphology, 476 identification, 482, 488 isolation procedures, 480–481 Mueller tellurite medium, 340 PGT medium, 342 specimen collection, transport, and handling, 300–301, 480 Tinsdale agar, 345 toxin, 480, 488, 496 toxoid, 496 typing systems, 496 Corynebacterium diphtheriae biotype belfanti, 482, 488 Corynebacterium diphtheriae biotype gravis, 482, 486, 488 Corynebacterium diphtheriae biotype intermedius, 482, 488 Corynebacterium diphtheriae biotype mitis, 482, 488, 1327 Corynebacterium durum, 475, 478, 482, 488–490, 493–494 Corynebacterium falsenii, 482, 489 Corynebacterium freiburgense, 482, 489 Corynebacterium freneyi, 482, 484, 489, 494 Corynebacterium glucuronolyticum, 478–479, 481–482, 489–490, 492 Corynebacterium hansenii, 482, 484, 489, 494 Corynebacterium imitans, 482, 489–490 Corynebacterium jeikeium, 474, 478–480, 482, 490, 492, 497, 1184, 1188–1189, 1327, 1332 Corynebacterium kroppenstedtii, 474, 479, 482, 487, 490 Corynebacterium lipophiloflavum, 482, 490 Corynebacterium macginleyi, 478–479, 482, 484, 490, 1327 Corynebacterium massiliense, 482, 490 Corynebacterium mastitidis-like organism, 490, 496 Corynebacterium matruchotii, 475–476, 482, 489–490, 493–494 Corynebacterium matruchotii-like strain, 490– 491 Corynebacterium minutissimum, 474, 479, 482, 484, 487, 490–493 Corynebacterium mucifaciens, 482, 484, 486, 491, 493 Corynebacterium nigricans, 487; see also Corynebacterium aurimucosum Corynebacterium paurometabola, see Tsukamurella paurometabola Corynebacterium pilbarense, 483, 491 Corynebacterium propinquum, 483–484, 487, 491

n lxiii

Corynebacterium pseudodiphtheriticum, 479, 483–484, 491 Corynebacterium pseudogenitalium, 493 Corynebacterium pseudotuberculosis, 474, 479–480, 483–484, 488, 491, 493, 496 Corynebacterium pyruviciproducens, 483, 491–492 Corynebacterium resistens, 474, 479, 483, 492, 497, 1327 Corynebacterium riegelii, 479, 483, 492 Corynebacterium seminale, see Corynebacterium glucuronolyticum Corynebacterium simulans, 483, 492 Corynebacterium singulare, 483–484, 487, 491–492 Corynebacterium sputi, 483, 492 Corynebacterium stationis, 483, 492 Corynebacterium striatum, 474, 478–479, 483, 485–486, 492, 497, 1327 Corynebacterium sundsvallense, 483–484, 492–493 Corynebacterium thomssenii, 483–484, 493 Corynebacterium timonense, 483, 493 Corynebacterium tuberculostearicum, 474, 479, 483, 485, 490, 492, 496 Corynebacterium tuscaniense, 483, 485 Corynebacterium ulcerans, 474, 479, 480, 483–484, 488, 491, 493, 496 Corynebacterium urealyticum, 474, 478–481, 483, 490, 493, 496, 1327 Corynebacterium ureicelerivorans, 483, 491, 493 Corynebacterium xerosis, 483–485, 489, 493 Coryneform Gram-positive rods, 474–498 antimicrobial susceptibilities, 496–497 antimicrobial susceptibility testing, 1327–1328 clinical significance, 479–480 collection, transport, and storage of specimens, 480 description of genera, 474–478 direct examination, 480 epidemiology and transmission, 478–479 evaluation, interpretation, and reporting of results, 497 identification, 481–496 isolation procedures, 480–481 serologic tests, 496 taxonomy, 474–475 typing systems, 496 Coryza, 1408, 1771 Cough adenoviruses, 1771 adiaspiromycosis, 2115 Anisakis, 2493 arenaviruses, 1673–1674 Ascaris lumbricoides, 2451 Coccidioides, 2114 coronaviruses, 1569 dirofilariasis, 2499 Histoplasma capsulatum, 2114 influenza virus, 1471 Linguatula serrata, 2516 monkeypox virus, 1830 Paragonimus, 2487 parainfluenza virus, 1488 respiratory syncytial virus (RSV), 1500 Strongyloides stercoralis, 2457 Countercurrent immunoelectrophoresis, 1969, 1971, 2181 Coverslip correction collar, 9 Coverslips, 9 Cowan, S. T., 255

lxiv

n

SUBJECT INDEX

Cowpox virus clinical significance, 1831 epidemiology and transmission, 1828–1829 PCR assay, 1834–1835 serologic tests, 1836 Coxiella, 283 Coxiella burnetii, 222–223, 887, 1150–1155 antimicrobial susceptibilities, 1180, 1184, 1193, 1195 antimicrobial susceptibilities, treatment, and prevention, 1155 biothreat agent, 222–223 clinical significance, 1083, 1151–1153 collection, transport, and storage of specimens, 1153 description of, 1150 diagnostic tests, 1084 direct examination, 1153–1154 epidemiology and transmission, 222–223, 1083, 1150–1151 evaluation, interpretation, and reporting of results, 1155 identification, 1154 isolation procedures, 1154 nucleic acid detection, 1153–1154 serologic tests, 1084, 1154–1155 complement fixation, 1155 ELISA, 1155 IFA, 1154–1155 taxonomy, 1150 typing systems, 1154 Coxiella cheraxi, 1150 Coxiellaceae (family), 887, 1150 Coxsackie B virus, 1427 Coxsackieviruses, 1536–1537 CPC method, of digestion and decontamination, 560 CPE, see Cytopathic effect CPE ELISA system, 948 CRBSI (catheter-related bloodstream infections), interpretive criteria for, 19 Credentials, for molecular methods, 82 Creutzfeldt-Jakob disease (CJD), 1859–1864 decontamination and, 197, 206–207 genetic (gCJD), 1859–1862, 1864 iatrogenic (iCJD), 1859–1861 laboratory-acquired infections, 177–178 sporadic (sCJD), 1859–1864 variant (vCJD), 197, 206–207, 1859–1864 Crimean-Congo hemorrhagic fever virus, 1645, 1651, 2507 Crohn’s disease Mycobacterium avium complex, 541 Yersinia enterocolitica, 742 Yersinia pseudotuberculosis, 742 Cronobacter collection, transport, and storage of specimens, 722 description of genus, 715 epidemiology, transmission, and clinical significance, 719 identification, 723, 725 taxonomy, 714–715 Cronobacter condimenti, 716, 719 Cronobacter dublinensis, 716, 719 Cronobacter helveticus, see Franconibacter helveticus Cronobacter malonaticus, 716, 719, 725 Cronobacter muytjensii, 716, 719 Cronobacter pulveris, see Franconibacter pulveris

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 64

Cronobacter sakazakii, 31, 716, 719, 725 Cronobacter turicensis, 716, 719 Cronobacter universalis, 716, 719 Cronobacter zurichensis, see Siccibacter turicensis Croup adenoviruses, 1771 coronaviruses, 1569 human metapneumovirus, 1509 parainfluenza virus, 1488 viruses, specimens and methods for detection of, 1407 Cruciate, 1941 Crustacea (class), 2507, 2522 Cryoprotective agents for lyophilization, 165 for ultralow-temperature freezing, 163–164 Crypt a-Glo FL, comprehensive kit, 2441 Cryptic species, 1984 Crypto, 2441 Crypto Cel, 2295, 2441 Cryptobacterium, 920–921 Cryptobacterium curtum, 925–926, 930 Crypto-CELISA, 2295 Cryptococcal Antigen LA system, 1997 Cryptococcosis, 1971 clinical features, 1993–1994 specimens for, 1947, 1949 Cryptococcus, 1937, 1990 antibody detection, 1971 antifungal resistance, 2243–2244 antifungal susceptibilities, 2006, 2224 antifungal susceptibility testing, 2259, 2263 antigen detection, 1977, 1996–1997 clinical significance, 1993–1994 cycloheximide inhibition, 1951, 1955 description of agents, 1988–1990 epidemiology and transmission, 1992 evaluation, interpretation, and reporting of results, 2006 identification, 1998–2003 in tissue, 1996 India ink preparation, 1988, 1996 media, 1959–1961 media for culture, 1952 microscopy, 1966, 1969, 1998 specimen collection, transport, and processing, 1948–1949 staining, 1956–1958 taxonomy, 1985 Cryptococcus adeliensis, 1985, 1993 Cryptococcus albidus, 1985, 1989–1990, 2002 Cryptococcus antigen test, 1997 Cryptococcus curvatus, 1985 Cryptococcus flavescens, 1985 Cryptococcus gattii, 1977, 1990, 1992, 2115 antigen detection, 1996 clinical significance, 1993–1994 identification, 1999–2000, 2002 media, 1959 multilocus sequence typing (MLST), 2004 staining, 1957–1958 taxonomy, 1985 typing, 2004 Cryptococcus laurentii, 1985, 1989–1990 Cryptococcus luteolus, 1989 Cryptococcus neoformans, 2115 antifungal resistance, 2225, 2229, 2238, 2241–2243 antifungal susceptibilities, 2005 antifungal susceptibility testing, 2258–2260, 2262–2263, 2265–2266

antigen detection, 1996 clinical significance, 1993 cultural and biochemical characteristics, 1989 description of agents, 1988–1990 detection in blood, 21 endophthalmitis, 1949 epidemiology and transmission, 1992 evaluation, interpretation, and reporting of results, 2006 identification, 1999–2003 India ink preparation, 1988 in tissue, 1996 isolation, 1998 media, 1959–1961 microscopy, 1973, 1975 multilocus sequence typing (MLST), 2004 stains, 1957–1958, 1970 taxonomy, 1985 typing, 2004 Cryptococcus neoformans var. grubii, 1985, 1992–1993 Cryptococcus neoformans var. grubii (serotype A), 1959, 1977 Cryptococcus neoformans var. neoformans, 1977, 1985, 1992–1993 Cryptococcus neoformans var. neoformans (serotype D), 1960 Cryptococcus terreus, 1989, 2002 Cryptococcus uniguttulatus, 1985, 1989 Cryptococcus uzbekistanensis, 1993 Cryptocotyle, 2482 Crypto-Giardia, 2441 Crypto-Giardia Ag Rapid test, 2441 Crypto/Giardia Cel, 2441 Crypto-LA test, 1997 Cryptosporidiosis, 125, 2435–2442 Cryptosporidium, 2405, 2435–2442, 2441 antigen detection, 2439 clinical significance, 2438 foodborne transmission, 2438 prevention, 2438 collection, transport, and storage of specimens, 2438–2439 commercial diagnostic assays, 2441 commercial kits for immunodetection in stool samples, 2295 culture, 2441 description of the agent, 2435–2436 detection, 2319–2320, 2326, 2329–2331, 2411–2412 direct examination, 2439 disinfection, 192–193 epidemiology, transmission, and prevention, 2436–2438 anthroponotic versus zoonotic transmission, 2437 susceptible populations, 2437 waterborne transmission, 2437–2438 evaluation, interpretation, and reporting of results, 2442 isolation procedures, 2440 life cycle, 2436 microscopy, 2439–2440 nucleic acid detection, 2308 sputum specimen, 2305 stains for detection, 2312–2314 taxonomy, 2435 treatment, 2440, 2535–2536 typing systems, 2440 Cryptosporidium Ag, ELISA, 2441 Cryptosporidium Ag Rapid test, 2441 Cryptosporidium andersoni, 2435 Cryptosporidium baileyi, 2435

SUBJECT INDEX Cryptosporidium bovis, 2435 Cryptosporidium canis, 2435, 2436, 2440 Cryptosporidium cuniculus, 2435, 2436, 2438 Cryptosporidium fayeri, 2435 Cryptosporidium felis, 2435, 2436, 2440 Cryptosporidium fragile, 2435 Cryptosporidium galli, 2435 Cryptosporidium genotyping kit, 2441 Cryptosporidium hominis, 2435–2440 Cryptosporidium macropodum, 2435 Cryptosporidium meleagridis, 2435, 2438 Cryptosporidium molnari, 2435 Cryptosporidium muris, 2435, 2440 Cryptosporidium parvum, 2405, 2435–2440 disinfection, 192 treatment, 2530, 2536, 2543 Cryptosporidium parvum antibody, 2441 Cryptosporidium Quik Chek, 2441 Cryptosporidium Rapid test, 2441 Cryptosporidium ryanae, 2435 Cryptosporidium scophthalmi, 2435 Cryptosporidium scrofarum, 2435 Cryptosporidium serpentis, 2435 Cryptosporidium suis, 2435, 2440 Cryptosporidium tyzzeri, 2435 Cryptosporidium ubiquitum, 2435, 2436 Cryptosporidium varanii, 2435 Cryptosporidium wrairi, 2435 Cryptosporidium xiaoi, 2435 Cryptosporium II test, 2295 Crypto-Strip C-1005, 2441 Crysomya, 2519 Crystal AutoReader, 364 Crystal E/NF Pseudomonas, 778 Vibrionaceae, 768 Crystal identification system anaerobic Gram-negative rods, 977 Clostridium, 954 Enterococcus, 409 non-spore-forming, Gram-positive, anaerobic rods, 927 Rapid Gram-Positive identification system, 364, 409 Crystal Neisseria/Haemophilus ID system, 676 CSF, see Cerebrospinal fluid (CSF) specimen CSF shunt infections Bacillus circulans, 443 Paenibacillus glucanolyticus, 443 Paenibacillus turicensis, 443 CSP (cell surface protein gene) typing, for Aspergillus, 2043 CT pELISA, 1116 CTBA, 481 Ctenocephalides, 2507 Ctenocephalides canis, 2509 Ctenocephalides felis, 2509 CTX-M β-lactamases, 695, 1223–1225, 1383–1384 Cul-de-sac fluid specimen collection, transport, and storage guidelines, 277 Culex, 2507 Culicoides, 2507 Culture, see also Blood culture amebae, pathogenic and opportunistic free-living, 2393–2394 fungal, 1944, 1947–1953 intestinal coccidia, 2430 larval-stage nematodes, 2321–2323 Leishmania, 2361 parasitology, 2297–2300 methods, 2307 urogenital samples, 2327 quantitative, 292

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 65

Trichomonas vaginalis, 2414–2415 Trypanosoma brucei, 2367 Trypanosoma cruzi, 2365 urine, 303–305 Culture media, 323–348 additives, 347–348 bacteriological, 325–347 composition of, 323–325 for detection of emerging antibiotic-resistant pathogens, 324 preparation of, 325 Cuneiform, 1941 Cunninghamella, 2088, 2096 Cunninghamella bertholletiae, 2088, 2096, 2101 Cunninghamella blakesleeana, 2096 Cunninghamella echinulata, 2088, 2096 Cunninghamella elegans, 2096 Cunninghamellaceae (family), 2088, 2096 Cupixi virus, 1669, 1671 Cupriavidus, 615 clinical significance, 795 description of genus, 792 direct examination, 795 epidemiology and transmission, 793 identification, 797–801 taxonomy, 791 Cupriavidus basilensis, 795 Cupriavidus gilardii, 615, 630–633, 791, 795 Cupriavidus metallidurans, 795 Cupriavidus necator, 791 Cupriavidus pauculus, 630–631, 791, 795 Cupriavidus respiraculi, 630–631, 791, 795 Cupriavidus taiwanensis, 791 Cu-PVA, 2311–2312 Current Procedural Terminology coding, 82 Curtobacterium chemotaxonomic features, 475 description of genus, 478 identification, 438, 484, 495 Curved and spiral-shaped Gram-negative rods algorithms for identification of, 994–997 Borrelia, 1037–1049 Campylobacter and Arcobacter, 998–1007 Helicobacter, 1013–1024 Leptospira, 1028–1033 Treponema and Brachyspira, 1055–1075 Curvularia, 1949, 1968, 2153, 2155, 2159–2163, 2167, 2174 Curvularia australiensis, 2155 Curvularia geniculata, 1968, 2155, 2161, 2174, 2177–2179 Curvularia hawaiiensis, 2155, 2160 Curvularia lunata, 1968, 2155, 2160–2161, 2174–2175, 2178–2179, 2181 Curvularia spicifera, 2155 Cutaneous infection/lesions, see also Skin infection/lesions anthrax, 125, 444 Coccidioides, 2114 Corynebacterium ulcerans, 479 dermatophytoses, 2135–2136 Fusarium, 2067 Helcobacillus, 479 Histoplasma capsulatum, 2114 hyaline fungi, 2076 leishmaniasis, 2358–2359, 2361 Microascus cirrosus, 2075 mucormycosis, 2089 Mycobacterium abscessus, 598–599 Mycobacterium chelonae, 596, 598–599 Mycobacterium kansasii, 542 Mycobacterium marinum, 542

n lxv

Mycobacterium szulgai, 543 phaeohyphomycoses, 2162 schistosomiasis, 2332 viruses, specimens and methods for detection of, 1406 Cutaneous larva migrans, treatment of, 2529, 2531–2532 Cutaneous specimen fungi, 1945, 1947, 1949 parasitology, 2294, 2298 CVM transport medium, 1410 CXCR4 coreceptor, 1437–1448, 1920 Cycloguanil, 2552–2553 Cycloheximide, fungi inhibition by, 1949–1951 Cyclophilin A, 1599 Cyclophyllidea (order), 2291, 2473–2475, 2501 Cyclopiazonic acid, 2189, 2191 Cycloserine activity, 1360 adverse effects, 1360 for Mycobacterium infection, 1360 Cycloserine-cefoxitin-egg yolk-fructose agar Clostridium difficile agar, 335 Cyclospora, 2425–2431 antimicrobial susceptibilities, 1192 stains for detection, 2312, 2316 Cyclospora cayetanensis, 2425–2431, 2440 antigen detection, 2430 clinical significance, 2428–2429 collection, transport, and storage of specimens, 2429 culture, 2430 description of agents, 2426–2427 detection, 2319 direct examination, 2429–2430 epidemiology, transmission, and prevention, 2427–2428 evaluation, interpretation, and reporting of results, 2431 laboratory techniques and control interventions used in significant outbreaks, 126 life cycles, 2426–2427 microscopy, 2426–2427, 2429–2430 nucleic acid detection, 2430 serologic tests, 2430 stains for detection, 2312–2314 taxonomy, 2425 treatment, 2431, 2530 Cylindrocarpon, 2063 Cylindrocarpon cyanescens, 2063 Cylindrocarpon destructans, 2063 Cylindrocarpon lichenicola, 2060 CyMol, 1409 Cyphellophora, 2154, 2156 Cyphellophora laciniata, 2154, 2156, 2162 Cyphellophora pluriseptata, 2154, 2156–2157, 2162 Cysteine albumin broth, 335 Cysteine heart agar (cystine-glucose-blood agar), 335 Cysteine-peptone-liver-maltose (CPLM) medium, 2315 Cystic echinococcosis, 2296 Cystic fibrosis patients Achromobacter xylosoxidans, 840–841, 843 Acrophialophora fusispora, 2076 Burkholderia, 791, 793, 795, 797, 803–804 Cupriavidus, 795 Dialister, 974 Haemophilus influenzae, 669

lxvi

n

SUBJECT INDEX

Cystic fibrosis patients (continued) Inquilinus limosus, 823 Mycobacterium, 541, 543, 599 Pandoraea, 795 Prevotella, 973 Pseudomonas aeruginosa, 774, 777–778, 783–784 Ralstonia, 795 rhinoviruses, 1553 Rhizobium radiobacter, 825 Scedosporium, 1950 Segniliparus rugosus, 519 Selenomonas, 974 specimen collection, transport, and handling, 299 Stenotrophomonas maltophilia, 794–795 Streptococcus anginosus group, 386 Cystic hydatid disease, 2471, 2476–2477 Cysticercoids, 2475 Cysticercosis, 2471, 2474–2477 commercial kits for immunodetection of serum antibodies, 2296 treatment, 2531–2533 Cysticercus, 2328–2330, 2332, 2473–2477 Cystine tryptic agar, 335 Cystine-tellurite-blood agar, 335, 481 Cystitis etiologies, usual, 291 viruses, specimens and methods for detection of, 1408 Cystoisospora, 2312, 2425–2431 Cystoisospora belli, 2425–2431, 2440 antigen detection, 2430 clinical significance, 2428 collection, transport, and storage of specimens, 2429 culture, 2430 description of agents, 2425–2426 detection, 2317–2319 direct examination, 2429–2430 epidemiology, transmission, and prevention, 2427 evaluation, interpretation, and reporting of results, 2431 life cycles, 2425 microscopy, 2426–2427, 2429–2430 nucleic acid detection, 2430 serologic tests, 2430 stains for detection, 2312–2314 taxonomy, 2425 treatment, 2430–2431, 2530 Cystoisospora natalensis, 2425, 2428 Cytarabine (ara-C) polyomavirus, 1811 progressive multifocal leukoencephalopathy (PML), 1811 Cytokine response modifier B (CrmB), 1833 Cytokines, varicella-zoster virus-stimulated, 1712 Cytomegalovirus (CMV), 1718–1732 antigen detection, 1722–1724 antiviral resistance, 1895–1896, 1917 antiviral susceptibility testing, 1730–1731, 1916, 1918–1919 clinical significance, 1719–1720 fetus and newborn infant, 1719 immunocompetent host, 1719 immunocompromised host, 1719–1720 collection, transport, and storage of specimens, 1720–1722 collection and handling, 1406–1408, 1413 specimens for direct detection, 1720– 1721

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 66

specimens for measurement of cellmediated immunity, 1722 specimens for serologic testing, 1721 specimens for virus isolation, 1721 storage and processing, 1411–1412 cytopathic effect (CPE), 1727 description of agent, 1718 detection and identification methods, 1433 DFA and IFA reagents for the detection of, 1425 direct examination, 1722–1726 disease prognosis by molecular methods, 76 epidemiology and transmission, 1718–1719 evaluation, interpretation, and reporting of results, 1731–1732 histopathologic testing, 1722 immunofluorescence in H&V-Mix cells, 1429 isolation procedures, 1725, 1727–1728 cell cultures, 1725, 1727 conventional tube culture, 1727 spin amplification shell vial assay, 1727–1728 nucleic acid detection, 1411, 1723–1726 rapid cell culture, 1426 serologic tests, 1728–1730 CVM IgM antibody measurements, 1729–1730 enzyme immunoassays, 1728–1729 IgG avidity assay, 1730 immunofluorescence assays, 1728–1729 measurement of CMV-specific cellmediated immunity, 1730 specimens for, 1721 taxonomy, 1718 TORCH (toxoplasmosis, other, rubella, cytomegalovirus, and herpes simplex virus) panels, 1530 treatment and prevention, 1720 vaccines, 1720 Cytomegalovirus (genus), 1398, 1718 Cytopathic effect (CPE), 1422, 1426–1427, 1429 adenovirus, 1775–1776 arboviruses, 1652 BK polyomavirus (BK virus), 1810 coronaviruses, 1577 cytomegalovirus (CMV), 1727 enteroviruses, 1543–1544 herpes simplex virus (HSV), 1690, 1692 human metapneumovirus, 1511–1512 influenza viruses, 1479 JC polyomavirus (JC virus), 1810 measles virus, 1522–1523 orthopoxviruses, 1835–1836 parainfluenza virus, 1491 parechoviruses, 1543–1544 polyomaviruses, 1810 respiratory syncytial virus, 1502, 1505, 1511–1512 Rhinoviruses, 1430 rhinoviruses, 1556–1557 varicella-zoster virus, 1710 Cytopenia, human herpesvirus 8 (HHV-8), 1762–1763 Czapek-Dox agar, 1961 D3 DFA identification and typing kit, for HSV, 1692 D3 DFA Respiratory Virus Screening & ID kit, 1473

D3 DFA VZV detection kit, 1710 D3 FastPoint, for parainfluenza virus, 1489 D3Ultra DFA HMPV, 1503, 1511 D3Ultra DFA respiratory kit, 1503 Daclatasvir, for hepatitis C virus infection, 1601 Daclatasvir resistance, 1902–1903 DAEC, see Escherichia coli, diffusely adherent Dalbavancin, 1187–1189, 1198 Dalfopristin, 1189–1190 Dangerous Goods Regulations, 1416 Danoprevir resistance, 1902 D’Antoni’s iodine, 2313 Dapsone activity, 1360 adverse effects, 1360 for Mycobacterium infection, 1360 Daptomycin, 1187–1189, 1198, 1256, 1260 Daptomycin resistance, 1229 Dark quencher probes, 61–62 Dark-field microscopy, 9 Brachyspira, 1063 Leptospira, 1031–1032 Treponema pallidum, 296–297, 1055, 1062– 1063, 1072 Darunavir, for human immunodeficiency virus (HIV), 1871, 1875 Darunavir resistance, 1897–1898 Databases, surveillance, 151 Davaineidae (family), 2291 Davidsohn differential test, 1743 DDT (dichlorodiphenyltrichloroethane), 2513 Deafness arenaviruses, 1673 Orientia, 1124 Debaryomyces, 1938 Debaryomyces fabryi, 1986 Debaryomyces hansenii, 1985–1986, 2000 Debaryomyces subglobosus, 1986 Debridement, 2518 DEC, see Diethylcarbamazine Decapods, 2507 Decarboxylase test, 317 Decolonization, 188–189 Decontamination cleaning and, 189–190 definition, 189 Decontamination agents, 320 Decoy cells, 1806–1807 Decubitus ulcer specimen collection, transport, and storage guidelines, 274 Deep wound, specimen collection from, 271 Deer ticks, 2512 DEET (diethyltoluamide), 2506 Defined medium (14, 17) for pathogenic Naegleria species, 2315 Definitive subtyping, definition, 132 Dehydration Cystoisospora belli, 2428 rotaviruses, 1622 Dehydroemetine, for Entamoeba histolytica, 2405 Delafield’s hematoxylin stain, 2314, 2333, 2336 Delafloxacin, 1179 Delayed hypersensitivity, atypical measles syndrome and, 1520 Delftia, 615 characteristics of, 802 clinical significance, 795 description of genus, 792 epidemiology and transmission, 793

SUBJECT INDEX identification, 802 taxonomy, 792 Delftia acidovorans, 630–631, 792, 795, 802, 804 Delirium, human herpesvirus 6 (HHV-6) and, 1756 Deltacoronavirus (genus), 1565 Deltaretrovirus (genus), 1399, 1458 Deltavirus (genus), 1399–1401 Delusory parasitosis, 2521 DeMan, Rogosa, Sharpe (MRS) agar (Lactobacillus MRS agar), 335 Dematiaceous, 1941 Dematiaceous fungi, 1969 cycloheximide inhibition, 1955 microscopy, 1967 polymorphous hyphae, 1967 Demeclocycline, 1185, 1187, 1198 Demetria, 354 Demodectic mange, 2517 Demodex, 2330 Demodex folliculorum, 2517 Dengue fever, 1647 arthropod vectors, 2507 laboratory techniques and control interventions used in significant outbreaks, 126 laboratory tests suggested for, 125 specimen collection and handling, 1406 Dengue hemorrhagic fever, 1654 Dengue shock syndrome, 1654 Dengue virus, 1406, 1644–1645, 1647–1649, 1652–1655 Density gradient media, 1422 Densovirinae (subfamily), 1818 Dental caries Actinomyces, 922 Bifidobacterium, 925 lactobacilli, 924 non-spore-forming, anaerobic, Gram-positive rods, 923 Olsenella, 925 Propionibacterium acidifaciens, 924 Streptococcus mutans group, 387 Dental culture, 274 Dental equipment, disinfection of, 200–201 Dental implant-associated infections, Actinomyces and, 922 Dental infections Actinomyces, 922 anaerobic Gram-negative rods, 974 Fusobacterium, 973 Gram-positive anaerobic cocci (GPAC), 910–911 Prevotella, 972 specimen collection, transport, and storage guidelines, 274 Denticle, 1941 Deoxycholate, 317 Deoxycholate agar, 335 Deoxycholate-citrate agar, 335 Deoxycholate-citrate agar, Hynes, 335 Deoxycholate-citrate-lactose-sucrose agar, 335 Deoxycholate-citrate-lactose-sucrose agar, Hajna, 335 Deoxynivalenol, 2190 Department of Health and Human Services (HHS), biothreat agents and, 217– 218 Dependovirus (genus), 1398 Depoendoparvovirus (genus), 1818 Depth of field, 5 Depth of focus, 5

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 67

Dermabacter, 478 antimicrobial susceptibility testing, 1328 chemotaxonomic features, 475 clinical significance, 479 description of genus, 475, 477 identification, 438, 484, 494 Dermabacter hominis, 475–479, 480, 486, 494 Dermabacteraceae, 478 Dermacentor, 2507, 2514–2515 Dermacentor albipictus, 2515 Dermacentor andersoni, 2515–2516 Dermacentor marginatus, 2515 Dermacentor occidentalis, 2515 Dermacentor variabilis, 2512, 2515 Dermacoccaceae family antimicrobial susceptibilities, 371 clinical significance, 361 description of family, 356 direct examination, 361 epidemiology and transmission, 357 evaluation, interpretation, and reporting of results, 372 identification, 366–367 isolation procedures, 362 taxonomy, 354 Dermacoccus, 354, 356 Dermacoccus nishinomiyaensis, 356, 366–367 Dermapak, 2137 Dermaptera (order), 2522 Dermasel Selective Supplement, 2139 Dermatitis cockroaches, 2513 Dermatophilus congolensis, 514–515 dermestid beetles and, 2521 human T-cell lymphotropic viruses (HTLVs), 1461 Malassezia, 1994 Sporobolomyces, 1994 swimmer’s itch (schistosome), 2480, 2486 urticating caterpillar hairs, 2519 Dermatobia, 2330 Dermatobia hominis, 2517, 2519 Dermatophagoides farinae, 2517 Dermatophilus chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 514–515 description of genus, 506 identification, 438 isolation procedures, 521 morphologic characteristics, 507 taxonomy, 505 Dermatophilus congolensis, 512–515, 521 Dermatophyte test medium (DTM), 1952, 1961, 2139 Dermatophytes, 2128–2146 anatomic specificity, 2136 anthropophilic species, 2135 antifungal resistance, 2245 antifungal susceptibility testing, 2271 antimicrobial susceptibilities, 2145 characteristics, 2129–2134 clinical significance, 2135–2136 description of etiologic agents, 2141–2145 epidemiology and transmission, 2135 evaluation, interpretation, and reporting of laboratory results, 2146 geophilic species, 2135 identification, 2139–2141 colony characteristics, 2140, 2144 growth on BCPMSG, 2141 growth on polished rice grains, 2141 in vitro hair perforation test, 2140

n lxvii

microscopic morphology, 2140 molecular techniques, 2141 nutritional requirements, 2140 physiological tests, 2140–2141 sequence of procedures, 2145 temperature tolerance and temperature enhancement, 2141 urea hydrolysis, 2141 isolation, 2138–2139 laboratory testing of specimens, 2137–2139 microscopy, 1967, 2137–2143 nucleic acid detection, 2139 specimen collection, transport, and processing, 1944, 1947, 1953, 2136–2137 strain typing systems, 2145 taxonomy, 2128 zoophilic species, 2135 Dermatophytoids, 2129–2134, 2146 Dermatophytosis, 2128 Dermestids, 2521 Desferrioxamine susceptibility, 616 Desmospora chemotaxonomic and lysosome growth characteristics, 509 description of genus, 506 morphologic characteristics, 507 Desmospora activa, 514 Desulfomicrobiaceae (family), 969 Desulfomicrobium, 969 Desulfomicrobium orale, 969, 981 Desulfovibrio characteristics of genus, 970–971 clinical significance, 974 identification, 976–977, 981–982, 994, 997 isolation procedures, 976 taxonomy, 969 Desulfovibrio desulfuricans, 974, 982 Desulfovibrio fairfieldensis, 974, 982 Desulfovibrio intestinalis, 974 Desulfovibrio orale, 974 Desulfovibrio piger, 969, 974, 977, 982 Desulfovibrio vulgaris, 974, 982 Desulfovibrionaceae (family), 969 Determine Syphilis TP, 1069 Deuteromycota, 1937 Deutsches Institut für Normung (DIN), 1268–1269 DFA, see Direct fluorescent antibody (DFA) test dfr gene, 1327, 1385 DGM-21A medium, 2315 Dhori virus, 1645 DHSP gene, Pneumocystis, 2026 Diabetes-associated infection, Bacillus cereus, 443 Diabetic foot infection Anaerococcus, 911 Bacteroides, 971 Finegoldia magna, 911 non-spore-forming, anaerobic, Gram-positive rods, 923 Pseudomonas aeruginosa, 775 Diacetoxyscirpenol, 2190 Diagnosis, by molecular methods, 74–75 Dialister antimicrobial susceptibilities, 984 characteristics of genus, 970–971 clinical significance, 974 identification, 976–977, 981 taxonomy, 969 Dialister invisus, 969, 974–975, 981

lxviii

n

SUBJECT INDEX

Dialister micraerophilus, 969, 974, 981 Dialister pneumosintes, 969, 974, 981 Dialister propionicifaciens, 969, 974, 981 Dialister succinatiphilus, 969, 981 Dialysis-associated infection Asaia, 829 Bacillus circulans, 443 Bacillus licheniformis, 442 Bacteroides, 970 Elizabethkingia meningoseptica, 828 Methylobacterium, 830 Pasteurella, 655 Roseomonas, 830 vancomycin-resistant lactobacilli, 924 Vibrio fluvialis, 765 Diamond-Blackfan anemia, 1820 Diamond’s complete medium modified by Klass, 2315 Diamond’s Trypticase-yeast extract-maltose (TYM) complete medium, 2315 Diaporthales (order), 2153–2155, 2159, 2173–2174 Diarrhea adenoviruses, 1770, 1772 Aeromonas, 754, 1326 anaerobic Gram-negative rods, 972 Anaerobiospirillum, 974 Anisakis, 2493 Arcobacter, 1001 arenaviruses, 1673 Bacillus subtilis, 442 Balantidium coli, 2417 Blastocystis hominis, 2406 Brachyspira, 1055 Campylobacter, 994, 1000–1001 Capillaria philippinensis, 2497 carbapenems, 1177 cephalosporins, 1175 clavulanic acid, 1177 clindamycin, 1185 Clostridium difficile, 905, 944 Clostridium perfringens, 942–944 coronaviruses, 1569 Cryptosporidium, 2438 Cyclospora cayetanensis, 2428, 2431 Cystoisospora belli, 2428, 2430–2431 daptomycin, 1189 Dientamoeba fragilis, 2413 Diphyllobothrium latum, 2472 Dysgonomonas, 654–655 Ehrlichia chaffeensis, 1138 Entamoeba histolytica, 2402–2403 enteric adenoviruses, 1618, 1621 Escherichia coli, 688–697 etiologies, usual, 290 filoviruses, 1674 fosfomycin, 1196 Giardia duodenalis, 2411 Grimontia hollisae, 766 hookworm, 2456 human herpesvirus 6 (HHV-6), 1755–1756 human herpesvirus 8 (HHV-8), 1762 Hymenolepis nana, 2476 influenza virus, 1471 linezolid, 1191 macrolides, 1183 measles, 1520 metronidazole, 1194 microsporidia, 2210, 2213, 2216 Moellerella wisconsensis, 722 monobactams, 1176 noroviruses, 1622 parechovirus, 1541

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 68

penicillins, 1173 Plesiomonas shigelloides, 721, 1326 rotaviruses, 1620 Salmonella, 701 Sarcocystis, 2429, 2431 Shigella, 697–699 spider envenomation, 2520 Strongyloides stercoralis, 2457 sulbactam, 1178 Taenia saginata, 2473 telithromycin, 1185 tetracyclines, 1187 Trichinella, 2495 Tropheryma whipplei, 1160–1161 Vibrio cincinnatiensis, 766 Vibrio fluvialis, 765 Vibrio furnissii, 765 Vibrio metschnikovii, 766 Vibrio mimicus, 763, 765 Vibrio parahaemolyticus, 765 viruses, specimens and methods for detection of, 1406 Yersinia enterocolitica, 742 DIC, filoviruses and, 1674 Dichloran, 1950 Dichlorodiphenyltrichloroethane (DDT), 2513 Dichotomomyces, 2030 Diclazuril, for Cystoisospora belli, 2431 Dicloxacillin, 1171–1172, 1198 Dicrocoeliasis, 2481, 2487 Dicrocoeliidae (family), 2290, 2481, 2487 Dicrocoelioidea (superfamily), 2290 Dicrocoelium, 2481 Dictyoptera (order), 2522 Didanosine, for human immunodeficiency virus (HIV), 1870, 1872 Didanosine resistance, 1896–1898 Dientamoeba, 2400, 2408 Dientamoeba fragilis, 2399–2400, 2405, 2408–2410, 2412–2413 clinical significance, 2413 description, 2412 detection, 2320, 2321 direct detection, 2413 epidemiology, transmission, and prevention, 2412–2413 evaluation, interpretation, and reporting of results, 2413 microscopy, 2413 taxonomy, 2412 treatment, 2413, 2530, 2542–2543 Diethylamide, 2190 Diethylcarbamazine, 2305, 2534–2535 adverse effects, 2535 filarial nematodes, 2464–2465 indications for, 2535 Loa loa, 2467–2468 mechanism of action, 2534 pharmacokinetics, 2534–2535 spectrum of activity, 2535 Diethyltoluamide (DEET), 2506 Dietzia acid-fast stain, 321 antimicrobial susceptibilities, 527 chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 515 description of genus, 506 identification, 438 microscopy, 521 morphologic characteristics, 507 taxonomy, 504–505 Dietzia maris, 515

Dietzia schimae, 515 Differential agar for group D streptococci, 335 Diff-Quik, 1970, 1975, 2333, 2335 Difluoromethyornithine, for Trypanosoma brucei, 2367 Digene CT/GC dual ID HC2, 638 Digene Hybrid Capture II, 1112 Digenea (subclass), 2290 Digenean trematodes foodborne, 2484, 2487–2490 life cycles, 2479, 2483–2484 of circulatory system, 2479–2486 Digital imaging, 51 Digital PCR, 63, 73 Dihydroartemisinin, 2540 Dihydrofolate reductase, 1191, 1234 Dihydropteroic acid synthase, 1234 Dikarya (subphylum), 2087 Diloxanide furoate, 2405, 2541 Dilution susceptibility testing, 1254–1264 agar methods, 1254, 1257–1258 advantages and disadvantages, 1258 dilution of antimicrobial agents, 1254 incubation, 1257 inoculation procedures, 1257 interpretation and reporting of results, 1257–1258 preparation, supplementation, and storage of media, 1257 antimicrobial agents recommended for routine testing, 1255–1256 breakpoints, 1259–1261 broth macrodilution method, 1258, 1261– 1262 advantages and disadvantages, 1262 dilution of antimicrobial agents, 1258 incubation, 1261 inoculation procedures, 1261 interpretation and reporting of results, 1261–1262 molds, 2268, 2270 preparation, supplementation, and storage of media, 1258, 1261 yeasts, 2262 broth microdilution method, 1262–1263 ability to detect resistance, 1278–1280 advantages and disadvantages, 1263 automated, 1275–1280 breakpoint susceptibility tests, 1263 dermatophytes, 2271 dilution of antimicrobial agents, 1262 gradient diffusion method, 1263 incubation, 1262–1263 inoculation procedures, 1262 interpretation and reporting of results, 1263 manual, 1275 molds, 2268–2271 preparation, supplementation, and storage of media, 1262 resistance screens, 1263 semiautomated, 1275 yeasts, 2258–2264 international methods, 1268 interpretive criteria, 1259–1261 quality control, 1263–1264 batch and lot QC, 1264 frequency, 1264 MIC ranges, 1264 reference strains, 1263–1264 single-well broth dilution method for clindamycin resistance detection

SUBJECT INDEX Staphylococcus, 1267, 1296 Streptococcus, 1267, 1298 Dimargaritales (order), 2087 Dimethyl sulfoxide (DMSO), storage of microorganisms in, 164 Dimorphic fungi, 2109–2123 antifungal susceptibilities, 2121–2122 antigen detection, 2116 biosafety, 2117 clinical significance, 2114–2115 collection, transport, and storage of specimens, 1948–1949, 1951, 2115 culture for mold phase, 2117 culture for yeast phase, 2118 description of agents, 2110–2113 direct examination, 2115–2117 epidemiology and transmission, 2113–2114 evaluation, interpretation, and reporting of results, 2121–2123 identification, 2118–2119 isolation, 2117–2118 media for culture, 1952 microscopy, 2110–2113, 2115–2116 nucleic acid detection, 2116–2117 serologic tests, 2120–2121 taxonomy, 2109–2110 typing systems, 2119–2120 Dimorphic pathogens, 1935, 1938 DIN (Deutsches Institut für Normung), 1268–1269 Dioctophymatidae (family), 2289 Dioctyphyma renale, 2328, 2330 Diopter adjustment, microscope, 13 Diorchitrema, 2482 Dipetalonema, 2328 Diphenhydramine, 2515 Diphetarsone, for Dientamoeba fragilis, 2413 Diphtheria, 479–480, 488; see also Corynebacterium diphtheriae Diphtheria toxin, 480, 488, 496 Diphtheria virulence agar base with tellurite and diphtheria virulence supplement, 335 Diphyllobothriasis, 2507 Diphyllobothriidae (family), 2291, 2471, 2502 Diphyllobothrium, 2332, 2471–2472 arthropod vector, 2507 detection, 2320 treatment, 2533 Diphyllobothrium cordatum, 2472 Diphyllobothrium dalliae, 2472 Diphyllobothrium dendriticum, 2472 Diphyllobothrium lanceolatum, 2472 Diphyllobothrium latum, 2471–2473 clinical significance, 2472–2473 collection, transport, and storage of specimens, 2473 description, 2471–2472 direct examination, 2473 eggs, 2449 epidemiology, transmission, and prevention, 2471–2472 evaluation, interpretation, and reporting of results, 2473 microscopy, 2473 nucleic acid detection, 2473 taxonomy, 2471 treatment, 2473, 2531 Diphyllobothrium pacificum, 2472 Diphyllobothrium ursi, 2472 Diphyllobothrium yonagoensis, 2472 Diplococcus morbillorum, 422

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 69

Diplococcus rubeolae, 422 Diploda (class), 2520, 2522 Diplomonadida (order), 2287, 2409 Diplostomida (order), 2290 Diplostomidae (family), 2290 Diplostomoidea (superfamily), 2290 Dipodascus capitatus, 1984 Dipstick immunoassay (DIA), for respiratory syncytial virus, 1502, 1504 Diptera (order), 2505–2508, 2522 Dipylidiasis, 2501–2502 Dipylidiidae (family), 2291, 2501 Dipylidium caninum, 2501–2502 arthropod vector, 2507, 2510, 2513 clinical significance, 2501 description of agent, 2501 detection, 2320 direct examination by microscopy, 2500–2502 epidemiology, transmission, and prevention, 2501 serologic tests, 2502 treatment, 2502, 2531 Direct fluorescent antibody (DFA) test, 97 Burkholderia pseudomallei, 795–796 Chlamydiaceae, 1110, 1112–1113 Cryptosporidium, 2439 detection of Chlamydiae and viruses, 1423–1425 Francisella tularensis, 856, 858 Giardia duodenalis, 2411–2412 herpes simplex virus (HSV), 1691–1692 human metapneumovirus (HMPV), 1503, 1510–1512 influenza viruses, 1472–1473 parainfluenza virus, 1489, 1491 parasitology, 2295 rabies virus, 1637–1640 respiratory syncytial virus, 1501–1503, 1508, 1510–1512 transport medium for, 1409 Trichomonas vaginalis, 2415 varicella-zoster virus, 1706, 1707 Direct fluorescent-antibody test for T. pallidum (DFA-TP), 1061–1063, 1073 Direct fluorescent-antibody tissue test for T. pallidum (DFAT-TP), 1062–1063 Direct rapid immunohistochemistry test (DRIT), for rabies virus, 1637–1639 Direct-acting antiviral drugs (DAAs), for HCV, 1601–1602, 1610–1611, 1878 Directigen A+B, 1474 Directigen EZ Flu A+B, 1474 Directigen EZ RSV, 1504 Directigen RSV, 1504 Direct-Ready MRSA panel, 361 Dirithromycin, 1260 Dirofilaria, 2499, 2501 Dirofilaria immitis, 2499–2501 clinical significance, 2499 description of agent, 2499 detection, 2328, 2330 direct examination by microscopy, 2499–2500 epidemiology, transmission, and prevention, 2499 serologic tests, 2499 treatment, 2499 Dirofilaria repens, 2499, 2501 Dirofilaria striata, 2501 Dirofilaria subdermata, 2501 Dirofilaria tenuis, 2499–2501 Dirofilaria ursi, 2499, 2501 Dirofilariasis, 2330, 2499

n lxix

Disability-adjusted life years (DALYs) liver flukes, 2489 Paragonimus, 2487 Disaster preparedness, 167 Discitis, Kingella and, 655 Discosphaerina filvia, 2155 Discriminatory power definition, 132 of subtyping method, 134 Disease outbreak (or cluster), see Outbreak Disease prognosis, molecular methods for, 76 Disease surveillance, 120–121 Disinfection/disinfectants, 190–202 antimicrobial activities, 190–191 by heat versus immersion in germicides, 201–202 choosing, guidelines for, 201 commonly used disinfectants for devices, 191–195 alcohols, 194 chlorine, 193 formaldehyde, 192–193 glucoprotamine, 195 glutaraldehyde, 191–192 hydrogen peroxide, 194–195 metals, 195 OPA, 192 overview, 201 peracetic acid, 194 peroxygen compounds, 195 phenolics, 194–195 quaternary ammonium compounds, 195 definition, 189, 190 hand, 186–187 inactivation of pathogens, 196 mode of application, 196 Mycobacterium resistance to, 600 of floors and surfaces, 196–197 of medical devices, 197–201 classification of devices, 197–198 dental equipment, 200–201 endoscopes, 198–200 of skin, 17 presurgical skin, 187–188 principles, 190–191 resistance to, 191, 195–196 surgical hand, 187 Disk diffusion method, 1247, 1264–1269 advantages and disadvantages, 1266–1267 agar medium for, 1265–1266 antifungal susceptibility testing molds, 2272 yeasts, 2266 antimicrobial agents recommended for routine testing, 1255–1256 breakpoints, 1259–1261 disks, antimicrobial agent, 1265–1266 fastidious bacteria, 1315, 1317–1318 for aminoglycoside resistance detection in enterococci, 1288 for extended-spectrum β-lactamase (ESBL) production confirmation, 1266 for mupirocin resistance detection in staphylococci, 1297 incubation, 1266 inhibition zone diameter distributions, 1249 inoculation procedure, 1266 international methods, 1268–1269 interpretation and reporting of results, 1254, 1266 interpretive categories, 1249, 1253–1254 interpretive criteria, 1259–1261, 1265 overview, 1264

lxx

n

SUBJECT INDEX

Disk diffusion method (continued) quality control, 1267 frequency of testing, 1267 reference strains, 1267 special disk tests, 1267 zone-of-inhibition diameter ranges, 1267 selection of antibacterial agents for routine testing, 1249 semiautomated instrumentation, 1274–1275 zone-of-inhibition diameter, 1259–1261, 1265, 1267 Disseminated intravascular coagulation, filoviruses and, 1674 Distal paresthesias, in tick paralysis, 2516 Distilled water, storage of microorganisms in, 162, 166 Distoseptate, 1941 Dithiothreitol, 1950, 1956 DiversiLab system, 137, 2003 Diversity index, 134 Dizziness, arenaviruses and, 1674 DMAIC cycle (define, measure, analyze, improve, and control), 47 DMSO, storage of microorganisms in, 164 DNA isolation methods, 77–78 quantifying in a sample, 73 DNA chip, 70, 71 DNA fingerprinting, 136, 2043 DNA gyrase, 1179, 1361 mutations in, 1218, 1232, 1361 DNA methylation, human papillomavirus (HPV) and, 1795 DNA microarrays described, 144–145 for pathogen identification, 144 for virus identification, 144–145 influenza viruses, 1480–1481 pathogen discovery, 241–42 DNA polymerase, 57; see also PCR; Target amplification techniques cytomegalovirus, 1895–1896 hepatitis B virus, 1899–1900 herpes simplex virus (HSV), 1894–1895 inhibition of, 78 varicella-zoster virus, 1895 DNA probes, 54 DNA sequence analysis for identification, 39, 75–76; see also Sequencing arthropods, 2523 classification and identification of bacteria, 260 Gram-negative nonfermentative rods, 816 Legionella pneumophila, 898 Neisseria, 642 non-spore-forming, anaerobic, Grampositive rods, 929 non-spore-forming, Gram-positive, anaerobic rods, 929 yeasts, 2002–2003 DNA target sequencing, for identification of organisms, 39 DNA topoisomerases, 1179 DNA-DNA hybridization, for classification and identification of bacteria, 259– 260 dnaJ gene, Legionella pneumophila, 898 DNase B, 396 DNase test agar with toluidine blue, 335 DNase-SISPA, 242 Dobrava-Belgrade virus, 1660–1662, 1664 Docosanol, for herpesviruses, 1884, 1886

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 70

Dog bites, see also Wound infection Corynebacterium canis, 487 Corynebacterium freiburgense, 489 Neisseria weaveri, 646 Dog tapeworm, see Dipylidium caninum Dog tick, 2512 Dog-biting louse, 2510 Dolosicoccus clinical significance, 425 identification, 426, 428, 429 taxonomy, 423 Dolosicoccus paucivorans, 423, 425 Dolosigranulum clinical significance, 424 identification, 427, 428, 429 taxonomy, 422 Dolosigranulum pigrum, 424, 428, 430 Dolphins, lacaziosis in, 2197 Dolutegravir, for human immunodeficiency virus (HIV), 1872, 1878 Dolutegravir resistance, 1447, 1897, 1899 DON, 2189–2190, 2190, 2192 Donovan bodies, 722 Donovanosis, 718 Dorea, 921 Doripenem, 1176–1177, 1255, 1260 Dothideales (order), 2148, 2153, 2155, 2159, 2163, 2166, 2173–2174 DOTS (directly observed therapy, shortcourse), 536 Double disk synergy test for AmpC enzymes, 1300 Double immunodiffusion, 95 Double-disk distortion test for detection of metallo-β-lactamases, 1301 Doxycycline, 1185–1186, 1198 antimicrobial susceptibility testing, 1256, 1260 ehrlichiosis, 2521 Lyme disease, 2521 Mansonella perstans, 2468 Plasmodium, 2349 Doxycycline resistance Bacillus anthracis, 1325 Francisella tularensis, 1325 Yersinia pestis, 1325 Dracunculiasis, 2496–2497 Dracunculidae (family), 2289, 2496 Dracunculoidea (superfamily), 2289 Dracunculus medinensis, 2496–2497 arthropod vector, 2507, 2513 clinical significance, 2496 description of agent, 2496 direct examination by microscopy, 2494, 2496 epidemiology, transmission, and prevention, 2496 serologic tests, 2496 treatment, 2496–2497 Drainage specimen, fungi, 1945, 1947–1948 Dried blood spots human immunodeficiency virus testing, 1445–1446 measles virus, 1521 rubella virus, 1527 viruses, collection methods and processing of specimens, 1413 Drug fever, cephalosporins and, 1175 Drug rash with eosinophilia and systemic symptoms (DRESS), HHV-6, 1756 Drug-induced hypersensitivity syndrome (DIHS), HHV-6, 1756 Dry objectives, 8 Drying, for storage of microorganisms, 162

Dryspot Campylobacter test kit, 1005 Dubos broth (Dubos Tween albumin broth), 336 Dubos Tween-albumin, 1364 Duet stains, 1427 Dulbecco’s phosphate-buffered saline, 1422 Duodenal capsule technique, 2326 Duodenal content specimens, 2325–2326 Duodenal contents, 2325–2326 Dust mites, 2512, 2517 Duvenhage virus, 1633–1634 Dye uptake (DU) test, 1914, 1916 Dyes, 320, 321 Dysentery bacillary dysentery, 697–699 Balantidium coli, 2417 Entamoeba histolytica, 2403 Shigella, 697–699 Trichuris trichiura, 2459 Dysgonomonas antimicrobial susceptibilities, 662 clinical significance, 654–655 direct examination, 656 epidemiology and transmission, 654 identification, 659–660 isolation procedures, 656 taxonomy and description of, 653 Dysgonomonas capnocytophagoides, 653–654, 657, 659, 662 Dysgonomonas gadei, 653, 655, 659 Dysgonomonas hofstadii, 653, 655, 659 Dysgonomonas mossii, 653, 655, 659 Dysphagia, Corynebacterium diphtheriae and, 480 Dyspnea Ascaris lumbricoides, 2451 Paragonimus, 2487 Strongyloides stercoralis, 2457 Dysuria, herpes simplex virus (HSV), 1688 D-zone test in staphylococci, 1267, 1296–1297 in streptococci, 1267, 1297–1298, 1319– 1320 E. coli O157:H7 MUG agar, 336 EAEC, see Escherichia coli, enteroaggregative Ear infection, see also Otitis anaerobic Gram-negative rods, 972 Ignavigranum, 425 Kerstersia, 841 Ear specimen collection, transport, and handling, 274, 295 Earle’s balanced salt solution, 1422 EAST1 (enteroaggregative ST-like toxin), 690 Eastern equine encephalitis (EEE) virus, 223–224, 1644, 1646–1648, 1651 EASY artus CMV PCR, 1726 Easy stain, 2441 EasyChip HPV blot chip, 1791 Ebola virus, 1647, 1669–1681 antigen detection, 1676 antiviral susceptibilities, 1681 clinical significance, 1674 collection, transport, and storage of specimens, 1675 description, 1670, 1672 direct examination, 1675–1677 epidemiology and transmission, 1670, 1673 evaluation and interpretation of results, 1681–1682 identification of virus, 1680 isolation procedures, 1678

SUBJECT INDEX laboratory tests suggested for, 125 nucleic acid detection, 1677 serologic diagnosis, 1680–1681 taxonomy, 1670, 1672 Ebolavirus (genus), 1398, 1670 EBURST algorithm, 139 EBV, see Epstein-Barr virus EBV-encoded RNAs (EBERs), 1738, 1742 Echidnophaga gallinacea, 2509 Echinacea, for rhinoviruses, 1558 Echinocandin(s), 2227–2228, 2244; see also specific drugs antifungal susceptibility testing, 2255– 2273 Aspergillus, 2044–2045 Candida, 2004 dimorphic fungi, 2121–2122 drug interactions, 2227 Fusarium, 2069 hyaline fungi, 2077 mechanism of action, 2227 mucormycosis, 2089, 2097 pharmacokinetics, 2227 spectrum of activity, 2227 therapeutic drug monitoring, 2228 toxicity and adverse effects, 2228 Echinocandin resistance, 2227, 2237, 2239, 2244–2245 Candida, 1993 epidemiology, 2244 intrinsic/inherent, 2237, 2245 mechanisms, 2244–2246 adaptive responses, 2245 biofilms, 2245 Fks, 2244–2246 hot-spot polymorphisms, 2245 Echinochasmus, 2482 Echinochasmus japonicus, 2490 Echinococcosis, 2296 Echinococcus detection, 2328–2330, 2332 treatment, 2531 Echinococcus granulosus, 2471, 2476–2477 commercial kits for immunodetection of serum antibodies, 2296 detection, 2329, 2331 sputum specimen, 2305 treatment, 2529, 2532 Echinococcus multilocularis, 2471, 2476–2477 animal inoculation, 2307 commercial kits for immunodetection of serum antibodies, 2296 treatment, 2531 Echinoparyphium, 2482 Echinostoma, 2320 Echinostoma hortense, 2482, 2484 Echinostomatidae (family), 2290, 2482, 2490 Echinostomatoidea (superfamily), 2290 Echinostomiasis, 2482 Echinulate, 1941 Echovirus, 1536–1537 Echovirus 30, transport medium for, 1409 EcoFix, 2311 Ectoparasites, treatment for, 2534; see also specific organisms Ectopic pregnancy, Chlamydia trachomatis and, 1108 Ectromelia virus, 1402 Eczema vaccinatum, 1831 Edema Capillaria philippinensis, 2497 Linguatula serrata, 2516 Onchocerca volvulus, 2466–2467 scorpion venom, 2519

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 71

Trypanosoma cruzi, 2362 urticating caterpillar hairs, 2519 Edwardsiella, 721 Edwardsiella hoshinae, 718, 721 Edwardsiella ictaluri, 718, 721 Edwardsiella tarda, 718, 721–722, 727, 730 EEE (Eastern equine encephalitis) virus, 223–224, 1644, 1646–1648, 1651 Efavirenz, for human immunodeficiency virus (HIV), 1870, 1874 Efavirenz resistance, 1897–1898 Efflux pumps, 1218–1219 azole resistance, 2241 chloramphenicol resistance, 1228–1229 macrolide resistance, 1231 polymyxin resistance, 1232 Pseudomonas aeruginosa, 781 quinolone resistance, 1233 RND (resistance-nodulation-cell division) type, 1218–1220, 1234 tetracycline resistance, 1233–1234 tigecycline resistance, 1234 Eflornithine, 2544–2545, 2551, 2555, 2564 adverse effects, 2545 mechanism of action, 2544 pharmacokinetics, 2544–2545 spectrum of activity, 2545 Trypanosoma brucei, 2367 Egg identification, parasite, 2320 Egg yolk agar, 336 Eggerthella antimicrobial susceptibilities, 931 clinical significance, 925 identification, 930 taxonomy and description, 920–921 Eggerthella hongkongensis, 920 Eggerthella lenta, 920, 925–926, 930, 1343– 1344, 1381 Eggerthella sinensis, 920, 925, 930 Eggerthia catenaformis, 920–921, 930 EHEC, see Escherichia coli, enterohemorrhagic Ehrlich reagent, 318 Ehrlichia, 1135–1145 antimicrobial susceptibilities, 1144 clinical significance, 1083, 1138–1139 collection, transport, and storage of specimens, 283, 1139–1140 description of genus, 1135 diagnostic tests, 1085 epidemiology and transmission, 1083, 1136–1138 evaluation, interpretation, and reporting of results, 1144–1145 features of, 1137 laboratory confirmation, 1140–1142 phylogenetics, 1136 taxonomy, 1135 Ehrlichia canis, 1136–1137, 1139, 2513 Ehrlichia chaffeensis, 1130 antigen detection, 1140 antimicrobial susceptibilities, 1144 arthropod vector, 2507 biosafety, 1140 description of, 1135 diagnostic tests, 1085 direct examination, 1140 epidemiology and clinical diseases associated with, 1083 evaluation, interpretation, and reporting of results, 1144–1145 features of, 1137 human monocytic ehrlichiosis (HME), 1138–1142, 1144–1145

n lxxi

identification, 1142 isolation procedures, 1140–1141 laboratory confirmation, 1140–1142 microscopy, 1140–1141 nucleic acid detection, 1140 phylogenetics, 1136 serologic tests, 1085, 1142 Ehrlichia ewingii, 1136–1137, 1139, 1144– 1145 diagnostic tests, 1085 epidemiology and clinical diseases associated with, 1083 Ehrlichia muris, 1137 Ehrlichia muris-like agent, 1083, 1085, 1136– 1137, 1139, 1144–1145 Ehrlichia ruminantium, 1135–1137 Ehrlichiosis human granulocytic ehrlichiosis (HGE), 2507, 2521, 2523 human monocytic ehrlichiosis (HME), 1138–1142, 1144–1145 preexposure prophylaxis, 2521 EIA, see Enzyme immunoassay EIEC, see Escherichia coli, enteroinvasive 8-aminoquinolines, 2539 Eiken Latex test, 1997 Eikenella antimicrobial susceptibilities, 662 antimicrobial susceptibility testing, 1328 clinical significance, 655 direct examination, 656 epidemiology and transmission, 653 identification, 658, 660 isolation procedures, 656 serotyping, 661 taxonomy and description of, 653 Eikenella corrodens, 653, 655–658, 660–662, 895, 1328 Eimeriida (order), 2287, 2373, 2425, 2435 Eimeriorina (suborder), 2373 Ekbom syndrome, 2521 Elbivudine, for hepatitis B virus, 1881–1882, 1900 Elecsys HBeAg, 1847 Elecsys HBsAg, 1847 Elecsys HSV-1 IgG and HSV-2 IgG assays, 1693 Electron microscopy adenoviruses, 1770, 1773 arenaviruses, 1675 Aspergillus, 2042 astroviruses, 1619, 1623 coronaviruses, 1565, 1570 cytomegalovirus, 1718 Encephalitozoon bieneusi, 2212 enteric adenoviruses, 1619, 1623 Epstein-Barr virus (EBV), 1742 filoviruses, 1675–1676 gastroenteritis viruses, 1619, 1622–1623 hantaviruses, 1660–1661, 1663 hepatitis A virus, 1591 hepatitis C virus, 1602 hepatitis E virus, 1591 human bocavirus, 1823 human herpesvirus 8 (HHV-8), 1764 human metapneumovirus, 1510 immunoelectron microscopy, 1623, 1629 influenza viruses, 1472 noroviruses, 1619, 1623 parainfluenza virus, 1488 parvovirus B19, 1819, 1821 polyomaviruses, 1806–1808 poxviruses, 1832–1833 rabies virus, 1638

lxxii

n

SUBJECT INDEX

Electron microscopy (continued) respiratory syncytial virus, 1501 rotaviruses, 1619, 1623 sapoviruses, 1619, 1623 Tropheryma whipplei, 1162 varicella-zoster virus, 1705, 1707 Electrophoresis, 67; see also Pulsed-field gel electrophoresis Electrospray ionization (ESI), 72 Electrospray ionization mass spectrometry (ESI-MS) influenza viruses, 1481 Mycobacterium, 584 Staphylococcus, 365 Elek method, 488 Elementary body, 1106–1107, 1111–1112, 1115, 1135 Elephantiasis, 2462, 2464 ELISA, see Enzyme-linked immunosorbent assay ELISPOT, cytomegalovirus, 1722 ELITech ELITe MGB, 1379, 1381–1382 Elizabethkingia, 616, 813, 828 Elizabethkingia meningoseptica, 615, 813, 826– 829, 831 Elizabethkingia miricola, 624–625, 828 Ellinghausen and McCullough, modified, 329 Ellinghausen-McCullough-Johnson-Harris medium, 336 ELVIS (enzyme-linked virus-inducible systems), 1426–1427, 1429, 1692 ELVIS HSV test, 1693 ELVIS replacement medium, 1430 Elvitegravir, for human immunodeficiency virus (HIV), 1872, 1877–1878 Elvitegravir resistance, 1447, 1897, 1899 EMB agar, Levine, 336 EMB agar, modified, Holt-Harris and Teague, 336 embB gene, 1358–1359 EMEM pH 2-3, 1430 Emericella, 1937, 2030, 2040 Emericella nidulans, 1938, 2031–2032, 2035, 2041 Emericella quadrilineata, 2031–2033, 2035 Emericella rugulosa, 2031–2032 Emericella unguis, 2031–2032 Emericellopsis, 2071 E-Mix refeed medium, 1430 Emm genes, 396 Emmonsia, 2076 adiaspiromycosis clinical significance, 2115 epidemiology and transmission, 2114 evaluation, interpretation, and reporting of results, 2123 nucleic acid detection, 2117 collection, transport, and storage of specimens, 2115 description of agent, 2112–2113 identification, 2119 isolation, 2118 microscopy, 2112–2113, 2116 nucleic acid detection, 2117 taxonomy, 2110 typing, 2120 Emmonsia crescens, 2110, 2112–2113, 2117– 2120 Emmonsia parva, 2110, 2112–2113, 2118– 2120 Emmonsia pasteuriana, 2110, 2113, 2118– 2119 Empedobacter, 616, 813 Empedobacter brevis, 624–625, 826, 828

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 72

Empyema etiologies, usual, 290 Gemella, 424 Parvimonas micra, 911 Saccharomyces cerevisiae, 1994 Staphylococcus, 360 Emtricitabine hepatitis B virus, 1881, 1900 human immunodeficiency virus (HIV), 1870, 1872 Emtricitabine resistance, 1921 hepatitis B virus, 1851, 1900 HIV, 1896–1898 Enanthem, arenavirus, 1674 Encephalitis Acanthamoeba, 2390–2391, 2394 adenoviruses, 1773, 1778 arboviruses, 1647 arthropod vectors, 2507 Balamuthia mandrillaris, 2389, 2391 enterovirus, 1540 herpes B virus, 1697 herpes simplex virus (HSV), 1689–1690, 1696 human herpesvirus 6 (HHV-6), 1755– 1756, 1760 influenza virus, 1471 Japanese encephalitis virus, 1654 Listeria monocytogenes, 463 measles inclusion-body encephalitis (MIBE), 1521–1522 microsporidia, 2210, 2213 parechovirus, 1541 phaeohyphomycoses, 2163 rabies virus, 1635 Rickettsia, 1124 specimen selection, 1541 Toxoplasma gondii, 2375, 2381 Trypanosoma cruzi, 2362–2363 varicella-zoster virus, 1705, 1709 viruses, specimens and methods for detection of, 1407 Encephalitozoon, 2209–2212, 2214–2216 culture, 2307 detection, 2328–2330 Encephalitozoon cuniculi, 2209–2210, 2213, 2215–2216 Encephalitozoon hellem, 2209, 2213 Encephalitozoon intestinalis, 2210, 2213, 2215, 2305, 2330 Encephalomyelitis Borrelia, 1041 Burkholderia, 794 measles, 1520 Encephalomyocarditis, enterovirus, 1540 Encephalopathy Bartonella, 876 Borrelia, 1041 human herpesvirus 6 (HHV-6), 1756 human immunodeficiency virus, 1439– 1440 influenza virus, 1471 Loa loa, 2467 progressive multifocal leukoencephalopathy (PML), 1756 transmissible spongiform encephalopathies (TSEs), 1859–1864 Endemic infections, 112 Endo agar, 336 Endocarditis Abiotrophia, 424, 431 Aerococcus, 424 Aeromonas, 754 Aggregatibacter, 654

anaerobic Gram-negative rods, 970, 972 Arcobacter, 1001 Aspergillus, 1948, 2037 Bacillus cereus, 443 Bacillus circulans, 443 Bacillus licheniformis, 442 Bacillus subtilis, 442 Bacteroides, 970 Bartonella, 873–874, 876 Blastoschizomyces, 1992 Burkholderia, 794 Candida, 1948, 1993 Candida albicans, 1948 Capnocytophaga, 654 Chlamydia psittaci, 1109 Citrobacter, 720 Clostridium, 946 Comamonas testosteroni, 795 Coniochaeta mutabilis, 2076 Coprinus cinereus, 2075 Corynebacterium diphtheriae, 479, 480, 1327 Corynebacterium jeikeium, 479 Corynebacterium pseudodiphtheriticum, 479 Corynebacterium striatum, 1327 Corynebacterium timonense, 493 Corynebacterium tuberculostearicum, 479 Corynebacterium tuscaniense, 493 Coxiella burnetii, 1152 Delftia acidovorans, 795 Eikenella corrodens, 655 Engyodontium album, 2076 enterococcal, 1213 Finegoldia magna, 911 fungal, 1948 Fusobacterium, 973 Gemella, 424, 431 Gram-negative nonfermentative rods, 813 Gram-positive anaerobic cocci (GPAC), 910–911 Granulicatella, 424, 431 HACEK organisms, 653, 654, 655, 1328 Haemophilus parainfluenzae, 670 Histoplasma, 1948 hyaline fungi, 2075–2076 Kingella, 655 lactobacilli, 924 Lactococcus, 424 Leptotrichia, 974 MIC values reporting, 1254 Microascus, 2075 Micrococcaceae, 361 Moraxella, 814 Neisseria bacilliformis, 645 Neisseria elongata, 645 Neisseria flavescens, 645 Neisseria meningitidis, 637 Neisseria mucosa, 645 Neisseria sicca, 645 Neisseria subflava, 645 non-spore-forming, anaerobic, Grampositive rods, 923 Paenibacillus glucanolyticus, 443 Paenibacillus popilliae, 443 Pasteurella, 655 Peptostreptococcus anaerobius, 911 phaeohyphomycoses, 2163 Phialemonium obovatum, 2076 Propionibacterium acnes, 924 Pseudomonas, 776 Pseudomonas aeruginosa, 775 Ralstonia, 795 Raoultella terrigena, 718 Rothia, 479 Rothia mucilaginosa, 361

SUBJECT INDEX Scopulariopsis brevicaulis, 2075 Shuttleworthia satelles, 924 Staphylococcus, 360, 1230 Stenotrophomonas maltophilia, 794 Streptococcus bovis group, 387 Streptococcus mitis group, 386 Streptococcus salivarius group, 387 Tropheryma whipplei, 1159, 1161 Veillonella, 911 Weissella confusa, 424 Endoconidium, 1941 Endodontic infection Actinomyces, 922 anaerobic Gram-negative rods, 972 Centipeda periodontii, 974 Dialister, 974 non-spore-forming, anaerobic, Grampositive rods, 923 Olsenella, 925 Slackia exigua, 925 Endogonales (order), 2087 Endolimax, 2399–2400 Endolimax nana, 2321, 2399–2402, 2407– 2408 Endometrial tissue and secretions, 277 Endometritis Capnocytophaga, 654 Chlamydia trachomatis, 1108 etiologies, usual, 290 Gardnerella vaginalis, 479 Mycoplasma, 1092 Neisseria gonorrhoeae, 636 Ureaplasma, 1092 Endoparasitic arthropods, 2516–2518 Endophthalmitis Aspergillus, 1949, 2032 Bacillus cereus, 443 Bacillus circulans, 443 Bacillus laterosporus, 443 Bacillus licheniformis, 443 Blastomyces dermatitidis, 1949 Candida, 1949, 1993 Clostridium, 946 Coccidioides, 1949 Cryptococcus neoformans, 1949 etiologies, usual, 290 fungal, 1949 Fusarium, 2058, 2067 Histoplasma capsulatum, 1949 Leuconostoc, 424 Mycobacterium haemophilum, 542 Paenibacillus alvei, 443 Paracoccidioides brasiliensis, 1949 Pseudomonas, 776 Rothia mucilaginosa, 361 Sporobolomyces, 1994 Sporothrix schenckii, 1949 Endoreticulatus, 2209–2210, 2213 Endoscopes outbreaks associated with contaminated, 199 reprocessing, 198–200 Endoscopy Anisakis worm removal by, 2493, 2495 Giardia duodenalis, 2411 Endospore-forming bacteria, aerobic, 441– 456 antimicrobial susceptibilities, 455–456 clinical significance, 442–445 collection, transport, and storage of specimens, 445–447 description of genera, 441–442 direct examination, 448–450 epidemiology and transmission, 442

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 73

evaluation, interpretation, and reporting of results, 456 identification, 451–454 isolation procedures, 450–451 serologic tests, 454–455 taxonomy, 441 typing, 454 Endospores, 2205–2206 Endosymbionts Francisella-like, 851, 857 Wolbachia, 1135–1136, 1138–1139, 2465, 2467, 2468, 2535 Endotracheal aspirate screening specimens, 284 specimen collection, transport, and handling, 278, 298 Enflagellation experiment, 2393 Enfuvirtide, 1871, 1877, 1920 Enfuvirtide resistance, 1447, 1897, 1899 Engyodontium, 2064, 2073, 2076 Engyodontium album, 2064, 2076 Enoplida (order), 2289 Entamoeba, 166, 2399–2408 Entamoeba bangladeshi, 2399–2400, 2402, 2405 Entamoeba coli, 2317–2319, 2321, 2399– 2402, 2407–2408 Entamoeba dispar, 2399–2406 detection, 2320, 2324–2325 nucleic acid detection, 2308 Entamoeba gingivalis, 2399–2400, 2407 detection, 2329, 2331 sputum specimen, 2305 Entamoeba hartmanni, 2321, 2399–2400, 2402, 2407 Entamoeba histolytica, 2387, 2399–2406 antigen detection, 2404, 2406 clinical significance, 2403 commercial kits for immunodetection in stool samples, 2295 commercial kits for immunodetection of serum antibodies, 2296 culture, 2307 description, 2400–2402 detection, 2319–2321, 2324–2325, 2327– 2332, 2404, 2406, 2411–2412 direct examination, 2403–2405 epidemiology, transmission, and prevention, 2402–2403 evaluation, interpretation, and reporting of results, 2405–2406 media for culture, 2315–2316 microscopy, 2404 nucleic acid detection, 2308, 2405 serologic tests, 2405 sputum specimen, 2305 treatment, 1187, 1194, 2405, 2530, 2535, 2541–2544 trophozoites and cysts, 2401–2402 Entamoeba histolytica CELISA Path, 2295 Entamoeba histolytica II test, 2295 Entamoeba histolytica Quick Chek, 2404 Entamoeba moshkovskii, 2399–2400, 2402, 2405 Entamoeba polecki, 2321, 2399–2400, 2402, 2407 Entecavir, for hepatitis B virus, 1881–1882, 1900 Entecavir resistance, 1851, 1900, 1917 Enteric adenoviruses antigen detection, 1624–1625 cell culture, 1627 clinical significance, 1620 description of agents, 1619

n lxxiii

electron microscopy, 1619, 1623 epidemiology and transmission, 1620–1621 evaluation, interpretation, and reporting of results, 1628–1629 molecular detection assays, 1625–1627 PCR, 1626 taxonomy, 1618 typing systems, 1628 Enteritis adenoviruses, 1772 Arcobacter, 1001 Campylobacter, 1000–1001 Mycobacterium genavense, 542 Enteritis necroticans, 943, 948–949 Enteroactinococcus, 354 Enteroaggregative ST-like toxin (EAST1), 690 Enterobacter, 723 antibiotic resistance, 1214, 1217, 1224, 1227 antimicrobial susceptibilities, 727–730, 1174–1175, 1177–1178, 1193, 1195–1196 antimicrobial susceptibility testing, 1266 as ESKAPE pathogen, 714 β-lactamases, 1299 description of genus, 715 epidemiology, transmission, and clinical significance, 719 evaluation, interpretation, and reporting of results, 731 identification, 725 taxonomy, 714–715 Enterobacter aerogenes, 715–716, 719, 723, 1226 Enterobacter agglomerans, 719 Enterobacter amnigena, see Lelliottia amnigena Enterobacter arachidis, see Kosakonia arachidis Enterobacter asburiae, 715–716, 723 Enterobacter cancerogenus, 716, 723, 725 Enterobacter cloacae, 31, 719, 725 antibiotic resistance, 1218, 1226–1227 antimicrobial susceptibilities, 1174 β-lactamases, 1299 Enterobacter cloacae subsp. cloacae, 716, 723 Enterobacter cloacae subsp. dissolvens, 716, 723 Enterobacter cowanii, see Kosakonia cowanii Enterobacter gergoviae, see Pluralibacter gergoviae Enterobacter helveticus, see Franconibacter helveticus Enterobacter hormaechei, 719, 725 Enterobacter hormaechei subsp. hormaechei, 716, 723, 725 Enterobacter hormaechei subsp. oharae, 716, 725 Enterobacter hormaechei subsp. steigerwaltii, 716, 725 Enterobacter kobei, 716, 723 Enterobacter ludwigii, 716, 719, 725 Enterobacter nimipressuralis, see Lelliottia nimipressuralis Enterobacter oryzae, see Kosakonia oryzae Enterobacter oryzendophyticus, 716 Enterobacter oryziphilus, 716 Enterobacter pulveris, see Franconibacter pulveris Enterobacter pyrinus, see Pluralibacter pyrinus Enterobacter radicincitans, see Kosakonia radicincitans Enterobacter sakazakii, see Cronobacter Enterobacter turicensis, see Siccibacter turicensis

lxxiv

n

SUBJECT INDEX

Enterobacteriaceae (family), 685–687, 693, 699 antibiotic resistance, 1224–1225, 1227– 1228, 1233–1234, 1279 AmpC cephalosporinases, 728–730, 1299–1300 β-lactamases, 727–728, 1299–1302 carbapenemases, 728, 730, 1300–1303 extended-spectrum β-lactamase (ESBL), 324, 327, 722, 727–729, 1279, 1299 phenotypic methods for detecting, 1287, 1298–1302 antimicrobial susceptibilities, 695–696, 727–730, 1172–1178, 1180–1181, 1184–1187, 1190–1191, 1193–1195 antimicrobial susceptibility testing, 1250, 1254–1256, 1259–1261, 1270– 1271, 1279–1280 biochemical reactions, 687 blood culture, 18 collection, transport, and storage of specimens, 722 description of genera, 715 epidemiology, transmission, and clinical significance, 715–722 evaluation, interpretation, and reporting of results, 730–731 identification, 722–726 isolation procedures, 722 properties of, 763 taxonomy, 714–718 typing systems, 726–727 Enterobacterial repetitive intergenic consensus PCR (ERIC PCR), 605 Enterobius gregorii, 2453 Enterobius vermicularis, 2453–2454 clinical significance, 2453–2454 description, 2453 eggs, 2449–2450, 2453 larvae, 2453 worms, 2450, 2453 detection, 2320, 2323–2324, 2459 diagnosis, 2454 Dientamoeba transmission and, 2412 epidemiology and prevention, 2453 taxonomy, 2453 transmission and life cycle, 2453, 2454 treatment, 2454–2455, 2531–2532, 2534– 2535 Enteroblastic, 1941 Enterococci, see Enterococcus Enterococcosel agar, 336 Enterococcosel agar with vancomycin, 336 Enterococcosel broth, 336 Enterococcus, 355, 403–415 aminoglycoside resistance, 1278, 1286– 1289 agar dilution screening method for detecting, 1287–1288 broth microdilution screening method for detecting, 1288 disk diffusion screening method for detecting, 1288 antibiotic resistance, 1214, 1217, 1220– 1221, 1229–1230, 1234 aminoglycoside, 1278, 1286–1289 linezolid resistance, 1278 molecular detection, 1381–1382 phenotypic methods for detecting, 1286–1289 reporting, 1289 vancomycin resistance, 1278, 1288– 1289

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 74

antimicrobial susceptibilities, 413–415, 1172, 1174–1175, 1177–1191, 1179, 1181–1182, 1184–1185, 1193, 1196 antimicrobial susceptibility testing, 1249, 1254–1256, 1259–1261, 1267, 1270, 1277 β-lactamase, 1302 clinical significance, 405–407 commercial sources of chromogenic agar media for, 326 description of genus, 403–405 epidemiology and transmission, 405 evaluation, interpretation, and reporting of results, 415 identification, 426 by commercial systems, 409, 411 by conventional physiological testing, 409 by MALDI-TOF MS, 411 by molecular methods, 411–412 phenotypic characteristics used for, 410 isolation procedures, 407–409 microscopy, 407 nucleic acid detection, 407 phenotypic methods for detecting antibacterial resistance, 1286–1289 aminoglycoside resistance, 1286–1288 vancomycin resistance, 1288–1289 phylogenetic tree, 404 serological tests, 413 specimen collection, transport, and storage, 407 taxonomy, 403 typing systems, 412–413 vancomycin-resistant enterococci (VRE) antimicrobial susceptibilities, 413–415 collection, transport, and storage of specimens, 407 commercial sources of chromogenic agar media for, 327 detection, 407, 1288–1289, 1294 health care-associated, 406–407 isolation procedures, 408–409 media for detection, 324 molecular detection of antibacterial resistance, 1381–1382 rectal swab screening for, 303 reporting to infection prevention program, 112 surveillance cultures, 113–115 Trypticase soy agar, with sheep blood and vancomycin for, 346 Enterococcus alcedinis, 410 Enterococcus aquimarinus, 410 Enterococcus asini, 410 Enterococcus avium, 406, 409–410, 414 Enterococcus caccae, 406, 410 Enterococcus camelliae, 410 Enterococcus canintestini, 403–404, 406, 410 Enterococcus canis, 410 Enterococcus casseliflavus, 404–406, 409–411 antibiotic resistance, 1230, 1278, 1288– 1289 antimicrobial susceptibilities, 413–415 Enterococcus cecorum, 403–404, 406, 410 Enterococcus columbae, 403–404, 410 Enterococcus devriesei, 403–404, 410 Enterococcus diestrammenae, 409 Enterococcus dispar, 406, 410 Enterococcus durans, 403–406, 409–410, 414 Enterococcus eurekensis, 410 Enterococcus faecalis

antibiotic resistance, 1216–1218, 1221– 1222, 1228–1231, 1234, 1287– 1289 antimicrobial susceptibilities, 413–415, 1175, 1177, 1184, 1187, 1189– 1190, 1195, 1279 clinical significance, 405–406 description, 403–404 epidemiology and transmission, 405 identification, 409–412 isolation procedures, 408–409 molecular detection of antibiotic resistance, 1381 nucleic acid detection, 407 phenotypic characteristics, 410 potassium tellurite agar for, 342 reference strains, 1264, 1267, 1288–1289, 1295, 1297 taxonomy, 403 tetrazolium tolerance agar, 345 typing systems, 412 Enterococcus faecium antibiotic resistance, 1212, 1216–1218, 1221–1222, 1228–1231, 1286–1289 antimicrobial susceptibilities, 413–415, 1175, 1177, 1184, 1187, 1189– 1190, 1279 as ESKAPE pathogen, 714 clinical significance, 406 epidemiology and transmission, 405 identification, 409–412 isolation procedures, 408–409 molecular detection of antibiotic resistance, 1381 nucleic acid detection, 407 phenotypic characteristics, 410 taxonomy, 403 typing systems, 412 vancomycin-resistant detection in blood, 24 Enterococcus gallinarum, 404–406, 409–411, 413–415 antibiotic resistance, 1230, 1278, 1288– 1289 antimicrobial susceptibilities, 413–415 Enterococcus gilvus, 404, 406, 410 Enterococcus haemoperoxidus, 404, 410 Enterococcus hawaiiensis, 406, 409–410 Enterococcus hermanniensis, 410 Enterococcus hirae, 406, 410, 414, 1221 Enterococcus italicus, 406, 410 Enterococcus lactis, 410 Enterococcus lemanii, 410 Enterococcus malodoratus, 406, 410 Enterococcus moraviensis, 403–404, 410 Enterococcus mundtii, 404, 406, 410, 414 Enterococcus pallens, 403–404, 406, 410 Enterococcus phoeniculicola, 410 Enterococcus plantarum, 404, 410 Enterococcus pseudoavium, 406, 410 Enterococcus quebecensis, 410 Enterococcus raffinosus, 406, 409–410, 414 Enterococcus ratti, 409–410 Enterococcus rivorum, 410 Enterococcus rotai, 404, 410 Enterococcus saccharolyticus, 403–404, 410 Enterococcus silesiacus, 404, 410 Enterococcus solitarius, 422 Enterococcus sp. nov. CDC PNS-E3, 409 Enterococcus sulfureus, 404, 410 Enterococcus termitis, 404, 410 Enterococcus thailandicus, 406, 410, 414 Enterococcus ureasiticus, 410 Enterococcus ureilyticus, 404, 410

SUBJECT INDEX Enterococcus viikkiensis, 410 Enterococcus villorum, 409–410 Enterococcus xiangfangensis, 409 Enterocolitis, adenovirus, 1772 Enterocytozoon, 2209–2211, 2214 Enterocytozoon bieneusi, 2210, 2212–2216, 2328 Enterocytozoon intestinalis, 2210, 2212 Enteromonas, 2408 Enteromonas hominis, 2321, 2400, 2408–2410, 2416 Entero-Test, 2305, 2326 Enterotoxin Bacteroides fragilis, 970 Clostridium perfringens, 942–943 enterotoxigenic Escherichia coli (ETEC), 685–686, 688–690, 692–693, 695– 697 Staphylococcus, 360, 366 Enterovibrio, 762 Enterovirus (genus), 1551 Enteroviruses, 1536–1546 antigen detection, 1542 antiviral susceptibilities, 1545 cell lines, susceptible, 1543–1544 clinical significance, 1538–1541 cytopathic effect, 1543–1544 description of agents, 1536–1537 detection and identification methods, 1433 DFA and IFA reagents for the detection of, 1425 direct examination, 1542–1543 epidemiology and transmission, 1537–1538 evaluation, interpretation, and reporting of results, 1545–1546 genome organization, 1536–1537 identification, 1543–1544 isolation procedures, 1543 meningitis, laboratory tests suggested for, 125 nucleic acid detection, 1542–1543 rapid cell culture, 1426 serologic tests, 1545 specimen collection and handling, 1406– 1408, 1415 specimen storage and processing, 1411 taxonomic classification, 1395–1396, 1399 taxonomy, 1536–1537 typing systems, 1544–1545 Entomophthoraceae (family), 2087 Entomophthorales (order), 1936–1937, 2087, 2102–2103; see also Entomophthoromycosis Entomophthoromycetes (class), 2087, 2102 Entomophthoromycosis, 2087, 2098–2103 antifungal susceptibilities, 2103 clinical manifestations, 2099 collection, transport, and storage of specimens, 1947, 2099–2100 direct examination, 2100 epidemiology and transmission, 2098–2099 evaluation, interpretation, and reporting of results, 2103 identification, 2100–2103 isolation, 2100–2101 microscopy, 2100 nucleic acid detection, 2100 taxonomy, 2087 treatment and outcome, 2099 Entomophthoromycota (phylum), 2087, 2098, 2100, 2103 Entomophthoromycotina (subphylum), 1936–1937, 2087

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 75

Entomoplasma, 1089–1090 Entomoplasmataceae (family), 1089 Entomoplasmatales (order), 1089 Entry inhibitors, human immunodeficiency virus, 1440, 1447, 1871, 1877 Enuresis, Enterobius vermicularis, 2454 env gene/protein, HIV, 1436–1437, 1443 Envenomation, arthropod sting/bite, 2518– 2521 centipedes and millipedes, 2520 Hymenoptera, 2518 scorpions, 2519–2520 spiders, 2520–2521 urticating caterpillars, 2518–2519 Environment, screening cultures from, 114 Enzygnost Syphilis, 1068, 1071 Enzyme immunoassay (EIA), 98–100 adenoviruses, 1773–1774 antibody interference, 100 antiviral susceptibility testing, 1914, 1916 Aspergillus, 1969 Blastocystis hominis, 2406 Blastomyces, 1969 Blastomyces dermatitidis, 2116 Borrelia, 1044–1046 Brucella, 868 Campylobacter, 1002 Chlamydiaceae, 1110, 1113, 1116 Clostridium difficile, 951 Coccidioides, 1971, 1977, 2121 competitive, 98 coronaviruses, 1570 Cryptococcus, 1977 Cryptosporidium, 2439, 2441 cytomegalovirus (CMV), 1728–1729 defined, 92 Ehrlichia chaffeensis, 1142 Entamoeba histolytica, 2404, 2405 Epstein-Barr virus, 1739, 1743–1745 gastroenteritis viruses, 1623–1625 Giardia duodenalis, 2411–2412 hepatitis C virus, 1607–1608 herpes simplex virus (HSV), 1692, 1693– 1694, 1696 Histoplasma capsulatum, 1977, 2116 hook effect, 99–100 human immunodeficiency virus, 1444 human metapneumovirus, 1510 human T-cell lymphotropic viruses (HTLVs), 1462–1465 IgM measurement, 100 influenza viruses, 1472–1475, 1481 mumps virus, 1495 Mycoplasma, 1097–1098 Mycoplasma pneumoniae, 1097 noncompetitive, 98, 99 parainfluenza virus, 1492 parasitology, 2295–2296 plate variability, 99 polyomavirus, 1810 Pseudomonas, 780 quantification of, 95 respiratory syncytial virus, 1502, 1504– 1505, 1508, 1510 Rickettsia, 1128–1130 Strongyloides stercoralis, 2458 technical challenges, 98–99 Toxoplasma gondii, 2375–2379 Treponema, 1062, 1067–1068, 1070–1071, 1074 combined treponemal IgM/IgG EIAs, 1068 treponemal IgM or IgG EIAs, 1067 Trypanosoma cruzi, 2365

n lxxv

varicella-zoster virus, 1711 Enzyme-linked immunosorbent assay (ELISA) Anaplasma phagocyrophilum, 1143–1144 arboviruses, 1648, 1653–1654 arenaviruses, 1676, 1680 Aspergillus, 1972, 2038, 2043–2044 β-lactamase detection, 1383 Blastomyces, 1969 Borrelia, 1043 Brucella, 867–868 Candida, 1996 Clostridium perfringens, 948 coronaviruses, 1570, 1577 Coxiella burnetii, 1155 CT pELISA, 1116 defined, 92 dengue virus, 1648 Ehrlichia chaffeensis, 1142 enteroviruses, 1545 Epstein-Barr virus, 1744 eumycotic mycetoma, 2181 filoviruses, 1676, 1680 Francisella, 859 fungi, 1969, 1971 gastroenteritis viruses, 1628 hantaviruses, 1664–1665 Helicobacter, 1019, 1021–1023 hepatitis A virus, 1592 hepatitis E virus, 1592–1594 herpes B virus, 1697 herpes simplex virus (HSV), 1693–1694 Histoplasma capsulatum, 2116, 2120 human herpesvirus 6 (HHV-6), 1759 human herpesvirus 7 (HHV-7), 1762 human herpesvirus 8 (HHV-8), 1759, 1764 human metapneumovirus, 1512 human papillomavirus, 1792 Hymenolepis nana, 2476 influenza viruses, 1480 Lagenidium, 2204 Leishmania, 2361 liver trematodes, 2489 Loa loa, 2467 measles virus, 1524–1526 Mycoplasma pneumoniae, 1098 Onchocerca volvulus, 2466 Paragonimus, 2487 poxviruses, 1835–1837 Pseudomonas, 780 Pythium insidiosum, 2203 rhinoviruses, 1558 rubella virus, 1528–1529 schistosomes, 2486 Strongyloides stercoralis, 2458 Taenia saginata, 2474 Taenia solium, 2475 Talaromyces marneffei, 2048 Toxoplasma gondii, 2377 transmissible spongiform encephalopathies (TSEs), 1863 Treponema pallidum, 1067, 1075 Trypanosoma brucei, 2367 Trypanosoma cruzi, 2365 Wuchereria bancrofti, 2465 Enzyme-linked immunosorbent spot (ELISPOT), cytomegalovirus, 1722 Enzyme-linked virus-inducible systems (ELVIS), 1426–1427, 1429, 1692 Enzywell Syphilis Screen, 1068, 1071 EPA, disinfectants and, 191, 194, 196 Epaxal, 1585 EPEC, see Escherichia coli, enteropathogenic

lxxvi

n

SUBJECT INDEX

Epichloë, 2190 Epidemic(s) adenoviruses, 1771 arbovirus, 1644 influenza, 1470–1471 norovirus gastroenteritis, 1620, 1622 outbreak recognition and investigation, 112 West Nile virus, 1647 Epidemic curve, examples of, 123 Epidemic keratoconjunctivitis, adenovirus, 1772 Epidemic relapsing fever, 2507 Epidemic typhus arthropod vectors, 2507, 2511 biothreat agent, 223 transmission and disease, 223 Epidemiological concordance, definition, 132 Epidemiological cutoff values (ECV/ECOFF), 2044, 2255 Epidemiology, see Molecular epidemiology; specific organisms Epidermophyton, 1939, 1961, 2128, 2145 Epidermophyton floccosum, 2129, 2135–2136, 2143, 2145 Epidermophyton stockdaleae, 2145 Epididymitis etiologies, usual, 290 Stenotrophomonas maltophilia, 794 viruses, specimens and methods for detection of, 1408 Epididymo-orchitis, Mycoplasma and, 1091 Epifluorescence, 10 Epigastric pain hookworm, 2456 Trichuris trichiura, 2459 Epiglottitis Haemophilus influenzae, 669 specimen collection, transport, and handling, 301 Epileptic encephalopathy, human herpesvirus 6 (HHV-6) and, 1756 Episthmium, 2482 Epsilon toxin, 223 Epsilonproteobacteria (class), 1013 Epstein-Barr virus (EBV), 1738–1747 antigen detection, 1742 clinical significance, 1739–1740 commercial products for EBV diagnosis, 1744 description of the agent, 1738 detection and identification methods, 1433 development of antibodies to EBV antigens following primary infection, 1745 direct detection, 1742 epidemiology and transmission, 1738–1739 evaluation, interpretation, and reporting of results, 1746–1747 future perspectives, 1747 isolation and identification procedures, 1743 latency, 1738 microscopy, 1742 nuclear antigens, 1738–1739, 1742–1746 nucleic acid amplification techniques (NAATs), 1742–1743 serologic tests, 1743–1746 EBV-specific antibodies, 1743–1746 heterophile antibodies, 1743 specimen collection and handling, 1406– 1408, 1413, 1740–1741 taxonomy, 1738

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 76

viral load, 1740, 1742–1743, 1746–1747 EraGen MultiCode CMV primers, 1726 EraGen MultiCode-RTx HSV 1 & 2 kit, 1691 Eravacycline, 1186–1187 ERG genes azole resistance and, 2238–2242 polyene resistance and, 2243 Ergine, 2190 Ergonomics, microscopy and, 12–13 Ergonovine, 2190 Ergot alkaloids, 2189–2191 Ergotamine, 2189–2190 Ergotism, 2190 Ergovaline, 2190 Eristalis tenax, 2517, 2519 erm genes, 595, 1231, 1267, 1296–1297, 1316, 1320, 1347, 1383 ermX methylase gene, 1327 Ertapenem, 1176–1177, 1198 anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1255, 1260 Bacteroides fragilis group susceptibility percentages, 1350 Erwinia, 719 Erwinia dissolvens, 719 Erwinia herbicola, 719 Erwinia milletiae, 719 Erwinia persicina, 725 Erwinia uredovora, 719 Erysipeloid, 468 Erysipelothrix, 467–469 antimicrobial susceptibilities, 469, 1184, 1197 clinical significance, 468 description, 467 direct examination, 468 epidemiology and transmission, 467–468 evaluation, interpretation, and reporting of results, 469 identification, 438, 468 isolation procedures, 468 serologic tests, 469 specimen collection, transport, and storage, 468 taxonomy, 467 typing systems, 468–469 Erysipelothrix inopinata, 467 Erysipelothrix rhusiopathiae, 467–469 antimicrobial susceptibilities, 469 antimicrobial susceptibility testing, 1317, 1328 clinical significance, 468 description, 467 direct examination, 468 epidemiology and transmission, 467–468 evaluation, interpretation, and reporting of results, 469 identification, 468 isolation procedures, 468 serologic tests, 469 specimen collection, transport, and storage, 468 taxonomy, 467 typing systems, 468–469 Erysipelothrix tonsillarum, 467–468 Erysipelotrichaceae (family), 922, 929 Erythema infectiosum, 1819, 1822 Erythema migrans, 1037, 1041, 1043 Erythema nodosum, Yersinia pseudotuberculosis, 742 Erythromycin, 1182–1183, 1198

antimicrobial susceptibility testing, 1255, 1260 Dientamoeba fragilis, 2413 Erythromycin resistance, 1231, 1316, 1320 Erythroparvovirus (genus), 1818 Erythrovirus (genus), 1398, 1818 ESBL, see Extended-spectrum β-lactamase Eschar Orientia, 1124 Rickettsia, 1124 Escherichia, 685–697 antimicrobial susceptibilities, 695–696 clinical significance, 686–690 collection, transport, and storage of specimens, 690 description of genus, 686 direct examination, 690–691 antigen detection, 690 microscopy, 690 nucleic acid detection, 690–691 epidemiology and transmission, 686 evaluation, interpretation, and reporting of results, 696–697 identification, 693–695 nucleic acid-based methods, 694 phenotypic, 693 serotyping, 693–694 virulence testing, 694–695 isolation procedures, 691–693 serologic tests, 695 taxonomy, 685 typing systems, 695 Escherichia albertii, 685–687, 693 Escherichia blattae, 685–687 Escherichia coli, 685–697 AmpC β-lactamases, 695–696, 1299 antibiotic resistance, 1212, 1214, 1216– 1219, 1224–1228, 1231–1233, 1235, 1383 antimicrobial susceptibilities, 695–696, 727, 729, 1174, 1177–1178, 1180, 1183, 1186, 1191–1192, 1195– 1196 antimicrobial susceptibility testing, 1279– 1280 attaching-and-effacing (A/E) lesion, 688 blood culture, 18 clinical significance, 686–690 CNF-producing, 690 collection, transport, and storage of specimens, 690 commercial sources of chromogenic agar media for, 326 cytolethal distending toxin-producing, 690 description of genus, 686 diarrheagenic antimicrobial susceptibilities, 696 clinical significance, 688–690 evaluation, interpretation, and reporting of results, 697 isolation procedures, 691–693 virulence testing, 694–695 diffusely adherent (DAEC), 688–689 direct examination, 690–691 antigen detection, 690 microscopy, 690 nucleic acid detection, 690–691 enteroaggregative (EAEC), 685, 688–690, 692–693, 695–697 enteroaggregative ST-like toxin (EAST1), 690 enterohemorrhagic (EHEC), 685–686 enteroinvasive (EIEC), 688–690, 692–697 enteropathogenic (EPEC), 685–686, 688– 690, 692–693, 695–697, 1192

SUBJECT INDEX enterotoxigenic (ETEC), 685–686, 688– 690, 692–693, 695–697, 1180 epidemiology and transmission, 686 evaluation, interpretation, and reporting of results, 696–697 extended-spectrum β-lactamases, 1299 extraintestinal antimicrobial susceptibilities, 695–696 clinical significance, 686–688 evaluation, interpretation, and reporting of results, 696 virulence testing, 694 identification, 693–695, 725 nucleic acid-based methods, 694 phenotypic, 693 serotyping, 693–694 virulence testing, 694–695 isolation procedures, 691–693 laboratory tests suggested for, 125 locus for enterocyte effacement (LEE) pathogenicity island, 686, 688–689, 694–695 meningitis/sepsis-associated (MNEC), 686, 688 O26:H11, 685, 688 O96:H19, 689 O157:H7, 685–686, 688–692, 694–697, 2389 clonality, 148 commercial sources of chromogenic agar media for, 326 specimen collection, transport, and storage guidelines, 275 O157:NM, 688 reference strains, 1264, 1267, 1300–1301 serologic tests, 695 Shiga toxin-producing (STEC), 148, 685, 688–692, 694, 696–697 Shigella compared to, 685, 697 taxonomy, 685 typing systems, 695 uropathogenic (UPEC), 685–686 Escherichia fergusonii, 685–687, 693 Escherichia hermannii, 685–687, 693 Escherichia vulneris, 685–687, 693 Esculin azide broth, 336 Esculin hydrolysis, 317 aerobic Gram-negative bacteria identification, 615 Clostridium identification, 954 Streptococcus, 395 eSensor, 72 eSensor HCV genotyping test, 1606 eSensor Respiratory Viral Panel (RVP), 1477 adenovirus, 1775 parainfluenza virus, 1490 rhinoviruses, 1554–1555 eSensor XT-8 Respiratory Viral Panel, 1506, 1511 ESKAPE pathogens, 714 Esophageal microbiome, 229 Esophagitis, HSV, 1689 ESP Myco medium, 336 ESP system, 21 Espline TP, 1069 E-swab, 48 ETEC, see Escherichia coli, enterotoxigenic Etest AmpC assay, 1300 anaerobic bacteria, 907 anaerobic Gram-negative rods, 984 antimicrobial susceptibility testing in anaerobes, 1344–1345

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 77

Aspergillus, 2045 Bacillus, 1326 Bacillus anthracis, 1325 Campylobacter, 1327 Corynebacterium, 497, 1328 described, 1263 fastidious bacteria, 1314 GRD, 1295 Haemophilus influenzae, 1322 Helicobacter, 1023 Helicobacter pylori, 1329 hVISA (heterogeneous vancomycinintermediate S. aureus), 1295 MBL strip, 1301 molds, 2272 Mycoplasma, 1099 Neisseria gonorrhoeae, 1323 Neisseria meningitidis, 1324 Streptococcus, 1320 Vibrio cholerae, 1331 VISA/VRSA testing, 1295 yeasts, 2267 Yersinia pestis, 1325 Ethambutol, for Mycobacterium infection, 1358–1359 Ethionamide activity, 1360 adverse effects, 1360 antimicrobial susceptibility testing, 1365– 1367 for Mycobacterium infection, 1358, 1360 Ethionamide resistance, 1358, 1360, 1367 Ethylene oxide gas, 204 ETI-AB-AUK Plus, 1847 ETI-AB-Corek Plus, 1848 ETI-AB-EBK Plus, 1848 ETI-EBK Plus, 1847 ETI-MAK-2 Plus DiaSorin, 1847 Etravirine, for human immunodeficiency virus (HIV), 1870, 1874 Etravirine resistance, 1897–1898 Euamoebida (order), 2287, 2387, 2399 Euascomycetes (class), 2174 Eubacteriaceae (family), 922, 929 Eubacterium antimicrobial susceptibilities, 931, 1189 clinical significance, 924–925 identification, 926, 930 taxonomy and description, 921–922 Eubacterium barkeri, 922 Eubacterium biforme, 922 Eubacterium brachy, 930 Eubacterium budayi, 922 Eubacterium callanderi, 922, 925 Eubacterium contortum, 922 Eubacterium cylindroides, 922 Eubacterium dolichum, 922 Eubacterium eligens, 922 Eubacterium hadrum, see Anaerostipes hadrus Eubacterium hallii, 922 Eubacterium infirmum, 922 Eubacterium lentum, see Eggerthella lenta Eubacterium limosum, 922, 930 Eubacterium minutum, 922, 930 Eubacterium moniliforme, 922 Eubacterium nitritogenes, 922 Eubacterium nodatum, 922, 924–926, 930 Eubacterium plautii, 922 Eubacterium ramulus, 922 Eubacterium rectale, 922, 930 Eubacterium saburreum, see Lachnoanaerobaculum saburreum Eubacterium saphenum, 922, 924, 930 Eubacterium siraeum, 922

n lxxvii

Eubacterium sulci, 922, 924, 930 Eubacterium tenue, 922, 925, 930 Eubacterium ventriosum, 922 Eubacterium yurii, 922, 924, 930 EUCAST, see European Committee on Antimicrobial Susceptibility Testing Eucestoda (subclass), 2471, 2473–2475 Euglenozoa (phylum), 2287 Eugonic agar, 336 Eugonic LT100 medium base without Tween 80, 336 Eumycotic mycetoma, 2173–2183 antigen detection, 2178 antimicrobial susceptibilities, 2181–2182 clinical significance, 2176–2177 collection, transport, and storage of specimens, 2177 colony morphology, 2179–2180 description of agents, 2173–2176 direct examination, 2177–2178 epidemiology and transmission, 2176 evaluation, interpretation, and reporting of results, 2183 identification, 2179–2181 molecular, 2180–2181 morphological, 2179–2180 isolation, 2178 microscopy, 2175, 2177–2178 nucleic acid detection, 2178 serologic tests, 2181 taxonomy, 2173–2176 typing systems, 2181 Euparpyhium, 2482 European Aspergillus PCR Initiative, 1979, 2039 European bat lyssavirus 1, 1633–1634 European bat lyssavirus 2, 1633–1634 European Centre for Disease Prevention and Control, 1269 European Committee on Antimicrobial Susceptibility Testing (EUCAST) antifungal susceptibility testing, 2255– 2265, 2267–2271 antimicrobial susceptibility testing, 108, 1321–1322, 1329, 1332 Aspergillus, 2044–2045 breakpoints, 1248 Campylobacter, 1007 clinical and bacteriological response rates, 1248 confirmatory and supplementary test use, 1250 methods advocated by, 1253, 1269– 1270 Pasteurella, 661 phenotypic methods for detecting resistance, 1291–1292, 1299 Pseudomonas, 783 selection of testing method, 1247 Staphylococcus, 369–370 website, 1249, 1254, 1281 European sheep tick, 2512 European Society for Clinical Microbiology and Infectious Diseases, 1269 Eurotiales (order), 1937 Eurotiomycetes (class), 1937–1938, 2109 Eurotium, 1937, 2030 Eurotium amstelodami, 2031–2032 Eurotium chevalieri, 2031–2032 Eurotium herbariorum, 2031–2032, 2036 Eurotium repens, 2031–2032 Eurotium rubrum, 2032 Euseptate, 1941

lxxviii n

SUBJECT INDEX

Euthambutol activity, 1359 adverse effects, 1359 antimicrobial susceptibility testing, 1365 Evans, Alfred, 246 Evans’ modified Tobie’s medium, 2315 Evolution of pathogens, 147 Evolution2, 1099 Evolution3, 1099 Ewingella, 722, 726 Ewingella americana, 718 Exanthematous rash adenovirus, 1773 measles, 1520 viruses, specimens and methods for detection of, 1406 ExaVir Load assay, human immunodeficiency virus, 1443 Excitation filters, 10–11 Exfoliative toxins, 360 Exiguobacterium description of genus, 477 epidemiology and transmission, 479 identification, 438, 495 taxonomy, 474–475 Exiguobacterium acetylicum, 477, 484, 495 Exiguobacterium aurantiacum, 477, 495 Exophiala, 2153–2154, 2157–2158, 2161, 2162, 2165, 2176 Exophiala asiatica, 2154, 2158 Exophiala attenuata, 2154, 2158 Exophiala bergeri, 2154, 2157 Exophiala dermatitidis, 2154, 2157, 2161– 2163, 2165, 2167, 2268–2269 Exophiala jeanselmei, 1968, 1973, 2154, 2157, 2161–2162, 2173–2176, 2178–2179, 2181–2182 Exophiala oligosperma, 2154, 2157, 2161–2162 Exophiala phaeomuriformis, 2154, 2158, 2163 Exophiala spinifera, 2154, 2158, 2161, 2163 Exophiala werneckii, 2147 Exophiala xenobiotica, 2154 Exserohilum, 1940, 2153, 2159, 2162, 2164 Exserohilum longirostratum, 2161 Exserohilum mcginnisii, 2161 Exserohilum rostratum, 1978, 2155, 2161, 2163–2164 Extended-spectrum β-lactamase (ESBL), 1172, 1173–1175, 1178, 1224–1225, 1254, 1270–1271, 1279 β-lactamase inhibitors, 1178 cephalosporins and, 1173–1175 commercial sources of chromogenic agar media for, 327 Enterobacteriaceae, 324, 327, 722, 727–729, 1299 Escherichia coli, 695–696, 1299 media for detection, 324 molecular detection, 1383 penicillins and, 1172 tests for, 1299 Extension oculomycosis, 1949 Extracellular polysaccharide production, by Streptococcus, 395 Exudate specimen, for fungi, 1945, 1947– 1948 Eye fatigue, microscopy and, 13 Eye infection, see also Ocular infection Bacillus cereus, 443 Corynebacterium macginleyi, 479, 490 Corynebacterium mastitidis-like organism, 490 Lactococcus, 424 Moraxella lacunata, 813–814

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 78

ophthalmomyiasis, 2518 parasitology, 2294, 2298 Propionibacterium propionicum, 924 Tropheryma whipplei, 1161 Eye specimen fungi, 1945, 1947, 1949–1950 parasitology, 2298, 2328, 2330 specimen collection, transport, and handling, 274, 295 viruses, 1413 Eyepiece, 9 EZ One, 1542 Facial edema Linguatula serrata, 2516 Trichinella, 2495 Facial palsy Borrelia, 1041 varicella-zoster virus, 1705, 1709 Facklamia clinical significance, 425 identification, 425, 426, 428, 429 taxonomy, 423 Facklamia hominis, 423, 428 Facklamia ignava, 423 Facklamia languida, 423, 427 Facklamia sourekii, 423 Facultative myiasis, 2516, 2519 Faecalibacterium, 921 Faecalibacterium prausnitzii, 231, 925 Failure mode effects analysis (FMEA), 170 Falciformispora senegalensis, 1968 Falciformispora tompkinsii, 1968 Falciformispora, 2176, 2178–2179 Falciformispora lignatilis, 2174 Falciformispora senegalensis, 2174, 2176, 2182 Falciformispora tompkinsii, 2174, 2176 Faldaprevir resistance, 1902 False negatives, 81, 92 False positives, 81, 92 Famciclovir antiviral susceptibility testing, 1916 herpes simplex virus (HSV), 1689, 1919 herpesviruses, 1884 varicella-zoster virus, 1706 Famciclovir resistance, 1917 herpes simplex virus (HSV), 1895 varicella-zoster virus, 1895 Fannia scalaris, 2519 Fansidar, 2540–2541 Far East scarlet-like fever, 743 Far Eastern tick-borne rickettsiosis, 1125 Fasciola, 2488, 2490 Fasciola gigantica, 2481, 2488–2489 Fasciola hepatica, 2481, 2488–2489 detection, 2320 eggs, 2449 treatment, 2531, 2533 Fascioliasis, 2481, 2488–2490 Fasciolidae (family), 2290, 2481–2482, 2487–2490, 2490 Fasciolopsiasis, 2482 Fasciolopsis, 2482 Fasciolopsis buski, 2320, 2449, 2484, 2488– 2490 Fast Track Referral Model System, Mycobacterium and, 587 FASTA format, 2523 Fastidious bacteria, antimicrobial susceptibility testing for, 1265, 1314– 1332 Fatal familial syndrome (FFI), 1859–1861 Fatigue Cyclospora cayetanensis, 2428

tick paralysis, 2516 FDA, see Food and Drug Administration Fecal leukocyte examinations, 301 Fecal specimen, see also Stool specimen for Helicobacter, 1018–1020, 1023 Gram stain and plating medium recommendations, 286 specimen collection, transport, and storage guidelines, 275, 281, 301–302 Federal Select Agent Program, 167, 219 Feeley-Gorman agar, 336 Feet, malodorous, 479 Fennellia, 2030 Fennellia flavipes, 2031, 2034 Fennellia nivea, 2031 Ferric ammonium citrate, 317 Ferric chloride, 318, 319 Festuclavine, 2190 Fetal death adenoviruses, 1773 Ljungan virus, 1541 Fetal hydrops, 1819–1820 Fetal infection, see also Congenital infection congenital varicella syndrome, 1412 cytomegalovirus, 1412, 1718–1719 parvovirus B19, 1412 viruses, specimens and methods for detection of, 1407 Fever adenoviruses, 1771, 1772 Anaplasma phagocyrophilum, 1139 arboviruses, 1647 arenaviruses, 1673–1674 Ascaris lumbricoides, 2451 Bartonella, 876–877 blackfly fever, 2515 Borrelia, 1041 Chlamydia psittaci, 1109 chloramphenicol, 1193 clindamycin, 1185 Coccidioides, 2114 coronaviruses, 1569 Corynebacterium diphtheriae, 480 Cyclospora cayetanensis, 2428 Cystoisospora belli, 2428 cytomegalovirus, 1718–1719 dirofilariasis, 2499 Ehrlichia chaffeensis, 1138 Epstein-Barr virus, 1739 filoviruses, 1674 herpes B virus, 1697 herpes simplex virus (HSV), 1688–1689 Histoplasma capsulatum, 2114 human herpesvirus 6 (HHV-6), 1754– 1756, 1761 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1762– 1763 influenza virus, 1471 Leptospira, 1030–1031 liver trematodes, 2489 macrolides, 1183 malaria, 2339 Mansonella, 2468 monkeypox virus, 1830 Mycoplasma, 1092 nitrofurantoin, 1196 Orientia, 1124 Paragonimus, 2487 Parastrongylus, 2498 parvovirus B19, 1819 polymyxins, 1193 polyomaviruses, 1804 respiratory syncytial virus (RSV), 1500

SUBJECT INDEX Rickettsia, 1124 rifampin, 1195 rubella, 1526 Sarcocystis, 2429 specimen selection, 1541 spider envenomation, 2520 sulfonamides, 1192 Talaromyces marneffei, 2046 Trichinella, 2495 Trypanosoma brucei, 2366 Trypanosoma cruzi, 2362 Trypanosoma evansi, 2368 Trypanosoma lewisi, 2368 vancomycin, 1189 varicella-zoster virus, 1704 yatapoxviruses, 1831 Fidaxomicin, 1183, 1348 Field diaphragm, 7 Field’s stain, 2333, 2335 Fifth disease, 1819, 1822 Filamentous fungi, see Molds Filarial dance sign, 2464 Filarial nematodes, 2461–2468 Loa loa, 2467–2468 lymphatic parasites, 2461–2465 Mansonella, 2468 Onchocerca volvulus, 2465–2467 Filariasis arthropod vectors, 2507 commercial kits for immunodetection of serum antibodies, 2296 detection, 2331 treatment, 2531–2532 Filariidae (family), 2461 Filarioidea (superfamily), 2289, 2461, 2465, 2467–2468 Fildes enrichment agar, 336 Filifactor, 921–922 Filifactor alocis, 922, 924, 926, 930 Filifactor villosus, 924 FilmArray, 24, 45, 60, 74, 392, 1383–1384 FilmArray blood culture identification (BCID) assay, 365, 1380–1382, 1380–1384 FilmArray gastrointestinal panel, 690, 1627 FilmArray respiratory panel, 1477, 1506, 1511, 1543 adenovirus, 1775 Haemophilus, 677 Mycoplasma, 1094 parainfluenza virus, 1490 rhinoviruses, 1554–1555 FilmArray respiratory panel pouch, 1576 Filobasidiales (order), 1937 Filobasidiella, 1937 Filobasidiella bacillispora, 1985 Filobasidiella neoformans, 1985 Filobasidium, 1937 Filobasidium uniguttulatum, 1985 Filoviridae (family), 1669–1682 animal inoculation, 1678 antigen detection, 1676 antiviral susceptibilities, 1681 cell culture, 1678 clinical significance, 1674 collection, transport, and storage of specimens, 1674–1675 postmortem specimens, 1675 safety and security, 1674–1675 shipping, 1675 specimen collection, 1675 description of agents, 1669, 1672 direct examination, 1675–1677 electron microscopy, 1675–1676

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 79

epidemiology and transmission, 1670–1673 evaluation and interpretation of results, 1681–1682 identification of virus, 1679–1680 immunofluorescence, 1679–1680 isolation procedures, 1677–1678 nucleic acid detection, 1677 serologic diagnosis, 1680–1681 ELISA, 1680 IFA test, 1680 neutralization tests, 1680 Western blotting, 1680 taxonomic classification, 1398, 1400 taxonomy, 1670, 1672 typing antisera, 1679 Filoviruses, 1433, 1647, 1669–1682 Filter paper/slant culture technique, 2323 Finafloxacin, 1179–1180 Fine needle aspirate specimen, for Mycobacterium, 547 Finegoldia, 909, 911 Finegoldia magna, 910–915 antimicrobial susceptibilities, 913, 1348 clinical significance, 911 direct examination, 912 epidemiology, 910 identification, 912–915 taxonomy, 910 Fire ants, 2518 Firmicutes (phylum), 474, 940, 967–969, 983 biochemical characteristics of human Eubacterium-like organisms, 930 clinical significance, 924–925 identification, 929 in colonic microbiome, 230 taxonomy and description of agents, 920– 922 FISH, see Fluorescence in situ hybridization Fish tapeworm, see Diphyllobothrium latum Fistula, Mycobacterium haemophilum, 542 5′ Exonuclease PCR (TaqMan) assays, 60–61 5-Fluorocytosine resistance to, 2006 yeast species, MICs for, 2005 510(k) submission, 1274 Fixatives, for parasitology, 2310–2312 Fks mechanism of echinocandin resistance, 2244–2246 Flaccid paralysis, arboviruses and, 1647 Flagellates, 2400, 2408–2416 antigen detection, 2408–2409 collection, transport, and storage of specimens, 2408 description of agents, 2408 direct examination, 2408–2409 epidemiology, transmission, and prevention, 2408 evaluation, interpretation, and reporting of results, 2409 key to identification of intestinal flagellates, 2322 microscopy, 2408 nonpathogenic, 2416 taxonomy, 2408 Flagellum stains, for aerobic Gram-negative bacteria identification, 615 Flash sterilization, 204 Flaviviridae (family), 1399–1401, 1599, 1644–1645, 1669 Flavivirus (genus), 1399, 1599, 1644–1645, 1647, 1649–1650, 1652, 1655 Flaviviruses arthropod vector, 2507 laboratory tests suggested for, 125

n lxxix

Flavobacteriaceae (family), 615, 652, 813, 831 Flavobacterium, 895 Flavobacterium mizutaii, 615 Flavobacterium multivorum, see Sphingobacterium multivorum Flavobacterium spiritivorum, see Sphingobacterium spiritivorum Flavobacterium yabuuchiae, see Sphingobacterium spiritivorum Flavonifractor, 921 Flavonifractor plautii, 922, 930 Flea-borne spotted fever, 1124, 1129 Fleas, 2509–2510, 2515–2516, 2522 Flesh flies, 2513, 2517 Fletcher medium, 336–337 Flexal virus, 1669, 1671 Flexirubin pigment production, for aerobic Gram-negative bacteria identification, 615 Flexispira rappini, 1013 Flexivirga, 354 FlexTrans viral transport medium, 1410 Flies key to, 2522 muscoid, 2513 myiasis, 2516–2519 as vectors, 2505–2506, 2508 Flinders Island spotted fever, 1125 Flinders Technology Associates, 165 Floccose, 1941 Floor cleaning and disinfection, 196–197 Flotation for cestodes, 2473, 2476 ova and parasite (O&P) examination, 2304–2305 Flow cytometry methods for antifungal susceptibility testing, 2265–2266 microparticle assays for Epstein-Barr virus, 1743–1744 Fluconazole, 2225 antifungal susceptibility testing, 2255– 2273 Balamuthia mandrillaris, 2395 Candida, 1993, 2004–2005, 2006 dermatophytes, 2145 dimorphic fungi, 2121–2122 eumycotic mycetoma fungi, 2181–2182 Leishmania, 2361 mucormycosis, 2089 Pythium insidiosum, 2203 Scopulariopsis brevicaulis, 2077 spectrum of activity, 2224–2225 Trypanosoma cruzi, 2365 yeast species, MICs for, 2005 Fluconazole resistance, 1993, 2225, 2238– 2242 Flucytosine, 2229–2230 antifungal susceptibility testing, 2255– 2273 eumycotic mycetoma fungi, 2181–2182 phaeohyphomycosis, 2167 spectrum of activity, 2229 Talaromyces marneffei, 2048 Flucytosine resistance, 2229, 2239, 2245– 2246 Fluids Gram stain and plating medium recommendations, 286 initial sample handling, 285 Flukes, see Trematodes Fluorescence in situ hybridization (FISH), 10 Pseudomonas, 776 Staphylococcus, 365

lxxx

n

SUBJECT INDEX

Fluorescence in situ hybridization (FISH) (continued) Tropheryma whipplei, 1162, 1164 Fluorescence microscopy, 10–11 light-emitting diode (LED) microscopes, 550–551 Fluorescence resonance energy transfer (FRET), 11, 57, 60–61 Fluorescence-activated cell sorter (FACS) analysis, VZV, 1712 Fluorescent antibody to membrane antibody (FAMA), VZV, 1710–1711 Fluorescent antibody virus neutralization (FAVN) test, rabies virus, 1641 Fluorescent treponemal antibody absorption (FTA-ABS), 1065–1066, 1070, 1074 Fluorescent-enzyme immunoassay (FEIA), fungi, 1969, 1971 Fluoribacter, 887 Fluorite objectives, 8 Fluorochrome, 10–11 5-Fluorocytosine resistance to, 2006 yeast species, MICs for, 2005 Fluorognost HIV-1 IFA, 1446–1447 Fluoroquinolone(s), 1178–1180 adverse effects, 1180 for Mycobacterium infection, 1358, 1361 mechanism of action, 1179 pharmacology, 1179 polyomavirus-associated nephropathy (PVAN), 1811 poultry use of, 1214 spectrum of activity, 1179–1180 Fluoroquinolone resistance, 1232–1233 adverse effects, 1361 alterations in target enzymes, 1232–1233 Bacillus anthracis, 1324 Brevibacterium, 1328 Campylobacter, 1326 Clostridium difficile, 1348 common associations of resistance mechanisms, 1215 Dermabacter, 1328 due to decreased intracellular accumulation, 1233 Escherichia coli, 1383 Haemophilus influenzae, 1321 molecular detection, 1383 Neisseria gonorrhoeae, 1323 Neisseria meningitidis, 1324 Streptococcus pneumoniae, 1319 FluoroType MRSA, 361 Foley catheter specimen collection, transport, and storage guidelines, 273 Folinic acid, 1191, 2381 Follicle mite, 2517 Follicular conjunctivitis, adenoviruses and, 1772 Folliculitis Aeromonas, 754 Malassezia, 1994 Pseudomonas aeruginosa, 775 Fomivirsen, for cytomegalovirus, 1720 Fonsecaea, 2153–2154, 2158, 2161, 2166, 2168 Fonsecaea compacta, 1967, 2158 Fonsecaea monophora, 2154, 2158, 2163, 2165, 2167 Fonsecaea multimorphosa, 2154, 2158 Fonsecaea nubica, 2154, 2158 Fonsecaea pedrosoi, 1967, 1975, 2154, 2157– 2158, 2163, 2166–2167 Fontana-Masson stain, 1957, 1965, 1970

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 80

Food and Drug Administration (FDA) antimicrobial susceptibility testing, 1249, 1274, 1314, 1316, 1319–1320, 1322 in anaerobic bacteria, 1345 mycobacteria, 1363, 1365–1366 antiseptics and, 183–184 arbovirus tests, 1655 disinfectants and, 188, 191–192, 195, 198–200 molecular detection of antibacterial resistance, 1379, 1381–1382, 1384– 1385 outbreak response, 121 reprocessed single-use devices, 205–206 tuberculocidal test, 191 viral specimen storage and processing, 1411 Food animals, antimicrobial use in, 1214 Food poisoning Clostridium perfringens type A, 942–943, 948 specimen collection, transport, and handling, 302–303 Staphylococcus, 360 Foodborne illness Arcobacter, 1001 Bacillus cereus, 443–445, 448 Bacillus pumilus, 443 Bacillus subtilis, 442 Campylobacter, 998–1000 Cryptosporidium, 2438 digenean trematodes, 2484, 2487–2490 Escherichia coli, 686 gastroenteritis viruses, 1620–1621, 1629 hepatitis A virus, 1587–1588 Listeria monocytogenes, 463–464 mycotoxins, 2190–2192 noroviruses, 1620–1621 outbreak investigation, 128 Salmonella, 701 Sarcocystis, 2427–2428 Toxoplasma gondii, 2373–2375 Vibrio mimicus, 765 Vibrio parahaemolyticus, 765 Foodborne Viruses in Europe (FBVE) network, 151 FoodNet, 128, 998, 1000 Food-safety biothreats, 224 Foot infection Anaerococcus, 911 Bacteroides, 971 Brevibacterium, 479 Corynebacterium confusum, 487 Corynebacterium simulans, 492 Finegoldia magna, 911 non-spore-forming, anaerobic, Grampositive rods, 923 Pseudomonas aeruginosa, 775 Foot spa disease, Mycobacterium and, 598 Foreign body infection Arthrobacter, 479 Brevibacterium, 479 Cellulosimicrobium, 479 Corynebacterium amycolatum, 479 Corynebacterium diphtheriae, 479 Corynebacterium jeikeium, 479 Corynebacterium striatum, 479 Microbacterium, 479 Micrococcaceae, 361 non-spore-forming, anaerobic, Grampositive rods, 923 Staphylococcus, 357, 360 Forensic molecular epidemiology, 148

Formaldehyde, 192–193 Formalin, 2310 cell culture preservation, 1423 stool specimen preservation, 2301–2303 Formicoidea, 2518 Forssman antigen, 1743 Fosamprenavir, for human immunodeficiency virus (HIV), 1871, 1875 Fosamprenavir resistance, 1897–1898 Foscarnet antiviral susceptibility testing, 1916 cytomegalovirus, 1720 Epstein-Barr virus, 1739 herpes B virus, 1697 herpes simplex virus (HSV), 1689, 1695 herpesviruses, 1883, 1885 human herpesvirus 6 (HHV-6), 1760 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1763 varicella-zoster virus, 1712 Foscarnet resistance, 1917 cytomegalovirus, 1730, 1895–1896 herpes simplex virus (HSV), 1894–1895 varicella-zoster virus, 1895 Fosfomycin, 1196 adverse effects, 1196 concentration in serum, 1199 mechanism of action, 1196 pharmacology, 1196 spectrum of activity, 1196 4 N (NNNN) medium, 2315 4-Aminoquinolines, 2536–2537 Fourier transform infrared (FTIR) spectroscopy, for bacterial identification, 263 Frambesia, 1061; see also Yaws Francisella, 851–859 antigen detection, 856 clinical significance, 854–855 collection, transport, and storage of specimens, 283, 855–856 description of agents, 852 differentiation from similar genera, 852 direct examination, 856–857 epidemiology and transmission, 852, 854 evaluation, interpretation, and reporting of results, 859 identification, 857–858 isolation procedures, 856–857 microscopy, 856 nucleic acid detection, 856–857 serologic tests, 859 taxonomy, 851 typing systems, 858–859 Francisella guangzhouensis, 851, 853–854 Francisella halioticida, 851–855 Francisella hispaniensis, 851–855 Francisella noatunensis, 851–855 Francisella novicida, 851–859 Francisella philomiragia, 851–856, 858–859 Francisella tularensis, 222, 851–859, 895 antigen detection, 856 antimicrobial susceptibilities, 1181, 1184 antimicrobial susceptibility testing, 1316, 1324–1325 arthropod vector, 2507 β-lactamase, 1325 biosafety, 855–856 biothreat agent, 220, 222 blood culture, 18 characteristics, 220 clinical significance, 854–855 collection, transport, and storage of specimens, 855–856

SUBJECT INDEX description of agents, 852 differentiation from similar genera, 852 direct examination, 856–857 epidemiology and transmission, 222, 852, 854 evaluation, interpretation, and reporting of results, 859 identification, 857–858 isolation procedures, 856–857 laboratory-acquired infections, 176 microscopy, 856 nucleic acid detection, 856–857 serologic tests, 859 taxonomy, 851 typing systems, 858–859 Francisella tularensis subsp. holarctica, 851– 856, 858–859 Francisella tularensis subsp. mediasiatica, 851– 855 Francisella tularensis subsp. tularensis, 851– 856, 858–859 Francisellaceae (family), 851, 857 Francisella-like endosymbionts, 851, 857 Franconibacter helveticus, 719 Franconibacter pulveris, 719 Freeze-drying (lyophilization), 164–165 cryoprotective agents, 165 fungi, 167 methods, 165 preparation of microbes, 165 reconstitution, 165 storage, 165 storage vials, 165 FreezerPro, 164 Freezerworks, 164 Freezing at –20°C, 162 fungi, 166 ultralow-temperature, 162–164 viral specimens, 1411 French Institute for Public Health Surveillance, 122 FRET (fluorescence resonance energy transfer), 11, 57, 60–61 Fretibacterium fastidiosum, 968, 974, 980–981 Fruiting bodies, 1936 FTA-ABS test, 1065–1066, 1070, 1074 FTD respiratory pathogens 21, 1555 FTD respiratory pathogens 33/21/21 plus, 1574 FTIR (Fourier transform infrared) spectroscopy, for bacterial identification, 263 Fugomyces cyanescens, 2071 Fumagillin, 2216 Fumigaclavines, 2190 Fumonisins, 2189–2191 Fungal meningitis, outbreak investigation, 127–128 Fungemia, 1948, 2059 Candida, 1993 detection, see Laboratory detection of bacteremia and fungemia Fusarium, 2058, 2067, 2068 hyaline fungi, 2075–2076, 2076 Pseudozyma, 1994 Saccharomyces cerevisiae, 1994 Fungi, see also Molds; Yeast(s); specific organisms biosafety, 1947 dimorphic, 1948–1949, 1951–1952, 2109– 2123 direct detection, 1965–1979 antibody detection, 1969, 1971

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 81

antigen detection, 1971–1972, 1975, 1977–1978 (1,3)-β-D-glucan detection, 1978 fungus-specific metabolite detection, 1978 mass spectrometry, 1978–1979 microscopy, 1965–1970, 1973–1976 nucleic acid detection, 1979 identification by nucleic acid sequencing, 75–76 of anamorphic molds, 1939–1940 of molds, 1940 of yeasts, 1939 polyphasic, 1940 key to medically important fungi, 1938 media, 1955–1956, 1959–1962 selection and incubation, 1951–1953 melanized, 2153–2168 microarrays for identification, 242 morphological characteristics, 1935–1936 mycotoxins, 2188–2192 nomenclature, 1936 reagents, 1956 risk-based classification, 171 saprophytic, 1951 specimen collection, transport, and processing, 1944–1953 specimen pretreatment, 1944, 1947–1948 specimen volume, 1944 stains, 1956–1959 storage methods, 166–167 taxonomy and classification, 1935–1940 terminology, 1940–1942 Fungi (kingdom), 1936–1938 Fungi Imperfecti, 1937 Fungitec, 1978 Fungitek G, 1996 Fungitell, 1978, 1996, 2024 Fungitest, 2265 Fungus balls, 2070, 2177 Fungus-specific metabolite detection, 1978 Funnel web spider, 2520 Furazolidone Giardia duodenalis, 2412 Trichomonas vaginalis, 2554 Furuncular myiasis, 2517 Furunculosis, Mycobacterium, 598 Fusariosis, 1947, 2057, 2068–2069 Fusarium, 1937, 1938, 1939, 1940, 2057– 2069, 2181 antifungal resistance, 2237, 2243–2244 antifungal susceptibility, 2069, 2224 antifungal susceptibility testing, 2268– 2269, 2271–2272 antigen detection, 2067–2068 classification of Fusarium infections, 2058 clinical significance, 2067 cycloheximide inhibition, 1955 description of agents, 2065 direct examination, 2067–2068 epidemiology and transmission, 2065, 2067 evaluation, interpretation, and reporting of results, 2069 identification, 2068 isolation, 2068 keratitis, 1949 key phenotypic features, 2060–2061 media, 1962 microscopy, 1967, 1969, 2067 nucleic acid detection, 2068 serologic tests, 2068–2069 specimen collection, transport, and processing, 1949, 2067

n lxxxi

taxonomy, 2059–2061, 2059–2065, 2065 trichothecenes, 2190 typing systems, 2068 Fusarium chlamydosporum, 2061, 2065 Fusarium chlamydosporum species complex (FCSC), 2061, 2065, 2069 Fusarium delphinoides, 2061, 2065, 2066 Fusarium dimerum, 2061, 2065, 2066 Fusarium dimerum species complex (FDSC), 2061, 2065–2066 Fusarium falciforme, 2060, 2061, 2065, 2067, 2173–2175, 2178–2180 Fusarium fujikuroi simplex complex (FFSC), 2060–2061, 2065, 2069 Fusarium incarnatum, 2061, 2065, 2069 Fusarium incarnatum-Fusarium equiseti species complex (FIESC), 2061, 2065, 2069 Fusarium keratoplasticum, 2060, 2061, 2065 Fusarium lichenicola, 2060 Fusarium MLST database, 2059 Fusarium moniliforme, 1967, 2060, 2066, 2069, 2261 Fusarium napiforme, 2060, 2061 Fusarium neocosmosporiellum, 2060, 2061 Fusarium nygamai, 2060, 2061 Fusarium oxysporum, 2065, 2067, 2069, 2271 Fusarium oxysporum species complex (FOSC), 1967, 2060–2061, 2065– 2067, 2069 Fusarium pallidoroseum, 2065 Fusarium petroliphilum, 2060, 2061, 2065 Fusarium poae, 2065 Fusarium proliferatum, 2060, 2061, 2065, 2066, 2067, 2069, 2190 Fusarium semitectum, 2061, 2065 Fusarium solani, 2067, 2069, 2261, 2271 Fusarium solani species complex, 1967 Fusarium solani species complex (FSSC), 2059–2061, 2065–2069 Fusarium solani var. coeruleum, 2177 Fusarium sporotrichioides, 2065 Fusarium thapsinum, 2060, 2065 Fusarium verticillioides, 2060, 2061, 2065, 2066, 2067, 2069 Fusarium-ID database, 2059 Fusidic acid, 1199 Fusiform, 1941 Fusion inhibitor(s), for HIV, 1440 Fusion inhibitor resistance, 1897, 1899 Fusobacteria (phylum), 967–968 Fusobacteriaceae (family), 967–968 Fusobacteriales (order), 967–968 Fusobacterium antibacterial resistance patterns, 1348 antimicrobial susceptibilities, 983–984, 1172, 1175, 1177, 1180, 1183– 1184, 1187, 1190, 1194, 1348, 1351 β-lactamase, 1348 characteristics of genus, 970–971 clinical significance, 973 direct examination, 975 epidemiology and transmission, 969 identification, 979, 981–983 isolation procedures, 976 taxonomy and description of genus, 968– 969 Vincent’s angina, 300 Fusobacterium alocis, see Filifactor alocis Fusobacterium canifelinum, 973 Fusobacterium fastidiosum, 981 Fusobacterium gonidiaformans, 968, 973, 981 Fusobacterium mortiferum, 968, 973, 977, 981, 1348

lxxxii n

SUBJECT INDEX

Fusobacterium naviforme, 968, 973, 979, 981 Fusobacterium necrogenes, 973 Fusobacterium necrophorum, 968, 973, 975– 977, 979, 981, 985 antimicrobial susceptibilities, 981, 1348, 1351 Lemierre’s disease, 905 specimen collection, transport, and handling, 299–300 Fusobacterium necrophorum subsp. animalis, 979 Fusobacterium necrophorum subsp. funduliforme, 973, 976, 979, 981 Fusobacterium necrophorum subsp. fusiforme, 979 Fusobacterium necrophorum subsp. necrophorum, 973, 976, 979, 981 Fusobacterium necrophorum subsp. nucleatum, 979, 981 Fusobacterium necrophorum subsp. polymorphum, 979 Fusobacterium necrophorum subsp. vincentii, 979 Fusobacterium nucleatum, 968–969, 973, 975– 977, 983, 1057, 1183, 1348, 1351 Fusobacterium perfoetens, 968 Fusobacterium periodonticum, 968, 973, 979, 981, 983 Fusobacterium russii, 968, 973, 979, 981 Fusobacterium simiae, 968, 973 Fusobacterium sulci, see Eubacterium sulci Fusobacterium ulcerans, 968, 973, 979, 981 Fusobacterium varium, 968, 973, 977, 979, 981, 1185 gag gene/protein, HIV, 1436–1437 Galactomannan, 1971–1972, 2038–2040 Gallibacterium anatis, 655 Gallicola, 909 Gallicola barnesae, 909 Gamma globulin, hepatitis A virus, 1590 Gamma interferon enzyme-linked immunospot (ELISPOT) assay, VZV, 1712 Gammacoronavirus (genus), 1565 Gammaherpesvirinae (subfamily), 1398, 1738, 1762 Gammapapillomavirus (genus), 1398, 1783 Ganciclovir adenoviruses, 1777 antiviral susceptibility testing, 1916 cytomegalovirus, 1720 Epstein-Barr virus, 1739, 1740 herpes B virus, 1697 herpesviruses, 1883, 1885 human herpesvirus 6 (HHV-6), 1760 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1763 Ganciclovir resistance, 1917 cytomegalovirus, 1730, 1895–1896, 1919 human herpesvirus 6 (HHV-6), 1760 Gardasil, 1786 Gardnerella antimicrobial susceptibilities, 498 chemotaxonomic features, 475 clinical significance, 479, 497 collection, transport, and storage of specimens, 497 direct examination, isolation, and identification, 497–498 epidemiology and transmission, 497 identification, 438 Gardnerella vaginalis, 474, 618, 2415 antimicrobial susceptibilities, 498

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 82

bacterial vaginosis, 479, 497–498, 925 clinical significance, 479, 497 detection, 2327 direct examination, isolation, and identification, 497–498 epidemiology and transmission, 497 Gram stain morphology, 476 identification, 484 specimen collection, transport, and handling, 296, 497 V agar for, 346 Gas gangrene, 906 Clostridium perfringens, 945–946 Clostridium septicum, 945 Clostridium sordellii, 946 collection, transport, and storage of clinical specimens, 292, 948 Gasterophilus, 2330 Gastric biopsy specimens, for Helicobacter, 276, 1018–1020 Gastric cancer, Helicobacter, 1017 Gastric lavage fluid specimen, for Mycobacterium, 276, 548 Gastric microbiome, 229–230 Gastritis etiologies, usual, 290 Helicobacter, 1017 Gastrodiscidae (family), 2482 Gastrodiscoides, 2482, 2489 Gastrodiscoidiasis, 2482 Gastrodiscus, 2482 Gastroenteritis, 1617 adenovirus, 1771–1772, 1778 Aeromonas, 754 Arcobacter, 996 astroviruses, 1618, 1621 Brachyspira, 996 caliciviruses, 1617, 1621 Campylobacter, 996, 1000 Cystoisospora belli, 2428 Edwardsiella, 721 enteric adenoviruses, 1618, 1621 Hafnia, 721 Helicobacter, 996, 1017–1018 Listeria monocytogenes, 463 noroviruses, 1617–1618, 1621 rotaviruses, 1617, 1620–1622 Salmonella, 701–702 sapoviruses, 1617, 1621 Tropheryma whipplei, 1161 Vibrio, 765–766, 996 Vibrio fluvialis, 765 Vibrio metoecus, 766 Vibrio mimicus, 765 Vibrio navarrensis, 766 viral, 1617–1629 Yersinia enterocolitica, 742, 747 Yersinia pseudotuberculosis, 742–743 Gastroenteritis viruses, 1617–1629 antigen detection, 1623–1625 antigenic and genetic typing systems, 1627–1628 cell culture, 1627 clinical significance, 1620–1622 collection, transport, and storage of specimens, 1622–1623 description of agents, 1618–1619 direct detection, 1623–1627 electron microscopy, 1619, 1623 epidemiology and transmission, 1619–1621 evaluation, interpretation, and reporting of results, 1628–1629 isolation procedures, 1627 molecular detection assays, 1625–1627

conventional PCR, 1625 electrophoresis, 1625 isothermal amplification assays, 1626– 1627 nanotechnology applied to testing, 1627 nucleic acid extract for testing, 1625 real-time PCR, 1625–1626 serologic tests, 1628 taxonomy, 1617–1618 Gastrointestinal disease Blastocystis hominis, 2406 coronaviruses, 1569 Entamoeba histolytica, 2402–2403 etiologies, usual, 290 influenza virus, 1471 Trichinella, 2495 Gastrointestinal microbiome, 229–231 Gastrointestinal tract specimen collection, transport, and handling, 271, 301– 303 beta-hemolytic streptococci, 303 Clostridium botulinum, 303 food poisoning, 302 Helicobacter pylori, 303 MRSA, 302–303 Shiga toxin-producing E. coli (STEC), 302 small bowel bacterial overgrowth syndrome, 303 viral, 1406 VRE, 303 Gastrospirillum hominis, see Helicobacter heilmannii Gatifloxacin, 1178–1180, 1199 Gbagroube virus, 1671 GBV-C, 1599 GC agar, 337, 1323, 1325 G+C content, 261 GC II agar, 337 GC-Lect agar, 337 Gel electrophoresis, 67 Gelatin hydrolysis aerobic Gram-negative bacteria identification, 615 Clostridium identification, 954 Gemella, 354 antimicrobial susceptibilities, 430 clinical significance, 424 description of genus, 423 epidemiology and transmission, 423 identification, 425, 427–428, 428, 429 interpretation of results, 431 taxonomy, 422 Gemella adiacens, 429 Gemella asaccharolytica, 422, 424, 428 Gemella bergeri, 422, 424, 428 Gemella haemolysans, 422, 423, 424, 427, 428, 429, 430 Gemella morbillorum, 422, 424, 428, 430, 927 Gemella sanguinis, 422, 423, 424, 427, 428 Gemifloxacin, 1178, 1199 Gen ID CAP Bac, 892 GEN III, 34, 453 GenBank, 2002 Gene expression profiling of pathogens, 71 Gene sequencing subtyping methods, see also specific methods CRISPR analysis, 140 multilocus sequence typing (MLST), 139 multivirulence locus sequence typing (MVLST), 139 whole-genome sequencing, 141 whole-genome SNP typing, 141–143 GeneFinder, 1791, 1795

SUBJECT INDEX GenElute, 929 GeneMark, 233 GeneOhm molecular detection of antibacterial resistance, 1379, 1381–1382 MRSA assay, 361, 1293 StaphSR assay, 1381 Staphylococcus, 365 VanR, 407, 1381 GeneSure MG test, 1922 Genetic reassortment, with influenza viruses, 1470 Genetic Systems HBsAg 3.0, 1847 Gen-Eti-K DEIA, 1606 GeneTrack HPV DNA chip, 1791 GeneXpert, 24, 45, 74 CT/NG real-time PCR assay, 1111 MTB/RIF assay, 1356, 1368 Staphylococcus, 365 Geniculate, 1941 Genital carcinoma specimens, 1408 Genital lesions adenoviruses, 1773 herpes simplex virus (HSV), 1688–1689, 1696 specimen collection, transport, and storage guidelines, 277 Genital tract infection Brucella, 865 etiologies, usual, 290 Eubacterium nodatum, 925 herpes simplex viruses, 1413 human papillomavirus (HPV), 1413–1414, 1784–1785 Mycobacterium genavense, 542 non-spore-forming, anaerobic, Grampositive rods, 923 Sneathia, 974 Genital tract specimens, 1408, 1413–1414 Genital ulcers Klebsiella pneumoniae subsp. granulomatis, 718 Treponema pallidum, 1058 Genitourinary infection adenoviruses, 1773 Alloscardovia omnicolens, 925 Burkholderia, 794 Corynebacterium aurimucosum, 479 Corynebacterium glucuronolyticum, 479 Gram-positive anaerobic cocci (GPAC), 910–911 Mycobacterium simiae, 543 Mycoplasma, 1091–1092, 1098 Peptostreptococcus anaerobius, 911 Roseomonas, 830 Ureaplasma, 1091–1092 Genitourinary schistosomiasis, 2480, 2484– 2485 GenoArray, 1791 GenoFlow, 1791 Genome sequence, see also Sequencing human microbiome, 232–233 Mycobacteria, 579 Tropheryma whipplei, 1159 GenoQuick MRSA, 361 Genotype 1+3 antigens, 1593 GenoType MRSA, 364, 366 GenoType MRSA Direct, 361 GenoType Mycobacterium CM/AS test, 582–583, 602 GenoType Staphylococcus, 365–366 Genotypic identification systems, 39–40 Genotypic resistance testing, VZV, 1712 Genotyping

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 83

Cryptosporidium, 2440–2441 human herpesvirus 8 (HHV-8), 1764 respiratory syncytial virus, 1505 rhinoviruses, 1557 GenPoint HPV detection system, 1789 GenProbe, 64 Mycoplasma genitalium, 1094 Pace 2, 638, 1112 Gentamicin, 1181–1182, 1199, 1255, 1260 Gentamicin resistance, in Mycobacterium, 1359 Gentamicin-amphotericin B solution (10X), 1423 Geobacillus, 441, 453 Geobacillus stearothermophilus, 441, 453 Geobacillus thermodenitrificans, 453 GeoSentinel, 128 Geosmithia argillacea, 2076 Geotrichum, 1960, 1991, 1998, 2001, 2263 Geotrichum candidum, 1987, 1989, 2004 Geotrichum capitatum, 1984 Germ tube test, 1999–2000 German cockroach, 2513 German Society for Hygiene and Microbiology, 1045–1046 Germicide, 195 definition, 190 disinfection by heat versus immersion in, 201–202 Gerstmann-Sträussler-Scheinker syndrome (GSS), 1859–1861 GG HPV chip, 1791 Gianotti-Crosti syndrome, human herpesvirus 6, 1755 Giant-cell pneumonia, measles, 1521–1522 Giardia, 2308, 2325–2326, 2408–2409, 2411– 2412, 2418 Giardia agilis, 2409 Giardia ardeae, 2409 Giardia duodenalis, 2399–2400, 2405, 2408– 2412 antigen detection, 2411–2412 clinical significance, 2411 description, 2409 detection, 2317–2319, 2322, 2325–2326 direct examination, 2411–2412 epidemiology, transmission, and prevention, 2409, 2411 evaluation, interpretation, and reporting of results, 2412 microscopy, 2411 nucleic acid detection, 2412 taxonomy, 2409 treatment, 2412, 2530, 2532, 2535–2536, 2542–2543 Giardia II, 2295 Giardia intestinalis, 2317–2319, 2322, 2325– 2326, 2409 Giardia lamblia, 2409 antimicrobial susceptibilities, 1194 commercial kits for immunodetection in stool samples, 2295 detection, 2317–2319, 2322, 2325–2326 Giardia microti, 2409 Giardia muris, 2409 Giardia psittaci, 2409 Giardia-Cel, 2295 Giardia-CELISA, 2295 Giardia/Cryptosporidium Chek, 2295, 2411 Giardia/Cryptosporidium Chek ELISA, 2441 Giardia/Cryptosporidium Quick Chek, 2295 Giardiasis, 125, 2294–2296, 2411–2412 Gibberella, 1937, 2059, 2189–2191

n lxxxiii

Gibberella fujikuroi, 2190 Gibberella nygamai, 2190 Gibberella pulicaris, 2189–2191 Gibberella tricincta, 2189–2191 Giemsa stain, 1957–1958 blood films for Plasmodium, 2341–2347 for blood parasites, 2334–2335 for parasitology, 2313–2314 fungi, 1970 Gigantobilharzia, 2480 Gimenez stain, 322 Gingivitis Peptostreptococcus anaerobius, 911 Prevotella, 972 Treponema, 1058, 1063 Gingivostomatitis, HSV, 1688 GLABRATA RTT, 2001 Glabrous, 1941 Glanders, 222, 794 Glarea lozoyensis, 2228 Glaucoma, rubella, 1526 Glimmer, 233 Global Microbial Identifier (GMI) network, 141 Global Outbreak Alert Response Network, 128 Global Polio Eradication Initiative, 1538 Global Public Health Intelligence Network, 128 Globicatella antimicrobial susceptibilities, 430 clinical significance, 425 identification, 426, 428, 429 taxonomy, 423 Globicatella sanguinis, 423, 430 Globicatella sulfidifaciens, 425, 428 Glomeromycota (phylum), 1936–1937, 2087 Glomerulonephritis measles, 1521 Streptococcus pyogenes, 385 Glossina, 2507 Gloves, 175–176, 283 Gloves and socks syndrome, 1819 Glucantime, 2542 Glucatell, 1978 Glucoprotamine, 195 Glutamate dehydrogenase, Clostridium difficile, 950–951 Glutaraldehyde, 191–192 Glycerol, storage of microorganisms in, 164 Glycine-buffered saline, 320 Glycopeptide(s), 1187–1189 adverse effects, 1189 antimicrobial susceptibility testing, 1260 mechanism of action, 1187–1188 pharmacology, 1188 spectrum of activity, 1188–1189 susceptibility reduction in staphylococci, 1279 Glycopeptide resistance, 1188, 1215, 1229– 1230 Glycylcyclines, 1185–1187 adverse effects, 1187 mechanism of action, 1186 pharmacology, 1186 spectrum of activity, 1186–1187 GN broth, Hajna, 337 Gnathostoma, 2497–2498 clinical significance, 2497–2498 description of agents, 2497 detection, 2328–2330, 2332 direct examination by microscopy, 2498 epidemiology, transmission, and prevention, 2497 serologic tests, 2498 treatment, 2498, 2531

lxxxiv

n

SUBJECT INDEX

Gnathostoma doloresi, 2497 Gnathostoma hispidum, 2497 Gnathostoma nipponicum, 2497 Gnathostoma spinigerum, 2497–2498, 2507, 2532, 2534 Gnathostomatidae (family), 2289, 2497 Gnathostomatoidea (superfamily), 2289 Gnathostomiasis, 2330, 2497–2498, 2507 Gnats, 2505 Golden Gate virus, 1672 Gomori methenamine silver (GMS) stain, for fungi, 1965, 1970, 1974, 1976 Gongylonema, 2513 Gongylonematidae (family), 2289 GonoCheck II, 641 Gonococcal Antimicrobial Surveillance Program (GASP), 1322–1323 Gonococcal Isolate Surveillance Program (GISP), 1322–1323 GonoGen II test, 641 Google Flu Trends, 128 Gordonia acid-fast stain, 321 chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 515 description of genus, 506 G+C content, 536 identification, 438, 523–524, 527 microscopy, 521 morphologic characteristics, 507 taxonomy, 504–505 Gordonia araii, 514–515 Gordonia bronchialis, 510–513, 515 Gordonia effusa, 514 Gordonia otitidis, 514–515 Gordonia polyisoprenivorans, 514, 527 Gordonia rubripertincta, 514 Gordonia sputi, 514 Gordonia terrae, 513, 515 Gordonibacter, 920–921 GP2, 453 Gradient diffusion method of antimicrobial susceptibility testing, 1263; see also Etest; M.I.C. Evaluator Gradient strip testing molds, 2272 yeasts, 2267 Graft-versus-host disease, human herpesvirus 6 (HHV-6) and, 1756 Grahamella, 873 Gram Positive 12 Easy-Plex PCR panel, 1381 Gram stain, 322; see also specific applications; specific organisms anaerobic bacteria identification and, 905–906 fungi, 1969, 1970, 1973, 1976 sampling handling, 285, 287 specimen types appropriate for, 285–287 Gram-negative anaerobic cocci (GNAC), 909–916 antimicrobial susceptibilities, 916 clinical significance, 911 description of group, 909 epidemiology, 910 evaluation, interpretation and reporting of results, 916 identification, 913, 916 isolation procedures, 912 taxonomy, 909 Gram-negative bacteria antibiotic resistance, 1217–1218, 1222– 1224, 1226, 1228–1228–1229, 1232–1234, 1280

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 84

β-lactamases in, 1383–1384 Gram-negative bacteria, aerobic dichotomous algorithms for identification bacteria with poor or no growth on sheep blood agar, 617 bacteria with purple or pink colonies on sheep blood agar, 618 glucose-fermenting bacteria (facultative anaerobes), 618 nonfermenters (strict aerobes), 619–623 nonfermenters, 613–634 schemes and tables, 616–634 identification, 613–634 identification tests, 613–616 acid production from carbohydrates/ sugars, 614 arginine dihydrolase, 614–615 assimilation/utilization of organic compounds as sole source of carbon, 614 colistin susceptibility, 616 desferrioxamine susceptibility, 616 esculin hydrolysis, 615 flagellum stains, 615 flexirubin pigment production, 615 gelatin hydrolysis, 615 growth at 41°C, 615 H2S production or sulfite reductase activity, 615 indole production, 615–616 motility, 616 NaCl requirement, 616 nitrite and nitrate reductase, 616 pyrrolidonyl aminopeptidase activity, 616 starch hydrolysis, 616 trypsin or benzyl-arginine arylamidase activity, 616 vancomycin susceptibility, 616 nonfermenters, 613–634 biochemical characteristics, 624–633 dichotomous algorithms for identification, 619–623 test methods, 613–616 Gram-negative rods anaerobic, 967–985 antimicrobial susceptibilities, 983–984 β-lactamase tests, 1302–1303 clinical significance, 969–974 collection, transport, and storage of specimens, 975 description of group, 967–969 direct examination, 975–976 epidemiology and transmission, 969 identification, 976–983 isolation procedures, 976 molecular detection, 975–976 reporting, interpretation, and reporting of results, 984–985 taxonomy, 967–969 unculturable, 983 antibacterial resistance patterns, 1348 curved and spiral-shaped algorithms for identification of, 994– 997 Borrelia, 1037–1049 Campylobacter and Arcobacter, 998–1007 Helicobacter, 1013–1024 Leptospira, 1028–1033 Treponema and Brachyspira, 1055–1075 fastidious or rarely encountered, 652–662 antimicrobial susceptibilities, 661–662 clinical significance, 654–655 collection, transport, and storage of specimens, 655

description of agents, 652–653 direct examination, 655–657 epidemiology and transmission, 653–654 evaluation, interpretation, and reporting of results, 662 identification, 658–661 isolation procedures, 656, 658 taxonomy, 652–653 typing systems and serologic tests, 661 nonfermentative, 813–831 antimicrobial susceptibilities, 830–831 clinical significance, 813–814 collection, transport, and storage of specimens, 814 description of agents, 813 direct examination, 814–815 epidemiology and transmission, 813 evaluation, interpretation, and reporting of results, 831 identification, 815–830 isolation procedures, 815 taxonomy, 813 Gram-positive anaerobic cocci (GPAC), 909–916 antimicrobial susceptibilities, 913, 916 clinical significance, 910–911 collection, transport, and storage of clinical specimens, 911–912 description of group, 909 direct examination, 912 epidemiology, 909–910 evaluation, interpretation and reporting of results, 916 identification, 912–915 differential characteristics, 914 flowchart for, 915 MALDI-TOF MS, 913 molecular methods, 913 isolation procedures, 912 taxonomy, 909–910 Gram-positive bacteria antibiotic resistance, 1217–1218, 1228– 1234, 1279 MALDI-TOF (MS) identification of, 37 Gram-positive cocci antibacterial resistance patterns, 1348– 1349, 1352 catalase-negative, 422–431 antimicrobial susceptibilities, 429–430 cellular morphology, 423 clinical significance, 424–425 collection, transport, and storage of specimens, 425 description of genera, 423 direct examination, 425 epidemiology and transmissions, 423– 424 evaluation, interpretation, and reporting of results, 430–431 identification, 425–429 isolation procedures, 425 serologic tests, 429 taxonomy, 422–423 typing systems, 429 catalase-positive, 354–372 antimicrobial susceptibilities, 368–371 clinical significance, 359–361 collection, transport, and storage of specimens, 361 description of families, 354–356 differentiating members of, 355, 357– 358 direct examination, 361–362 epidemiology and transmission, 356–357

SUBJECT INDEX evaluation, interpretation, and reporting of results, 371–372 identification, 362–367 isolation procedures, 362 serologic tests, 368 taxonomy, 354 typing systems, 367–368 identification of aerobic, 350–352 Gram-positive rods aerobic actinomycetes, 504–528 identification of, 437–439 antibacterial resistance patterns non-spore-forming bacilli, 1348 spore-forming bacilli, 1348 Clostridium, 940–959 coryneform, 474–498 identification, algorithm for, 437–439 non-spore-forming anaerobic, 920–932 antimicrobial susceptibilities, 931 clinical significance, 922–925 collection, transport, and storage of specimens, 925 direct examination, 925–926 epidemiology and transmission, 922 evaluation, interpretation, and reporting of results, 931–932 identification, 926–930 isolation procedures, 926 serologic tests, 931 taxonomy and description of agents, 920–922 Gram-Sure, 316 Granule cell neuronopathy, JC polyomavirus and, 1804 Granules, fungal, 1967–1968 Granulibacter bethesdensis collection, transport, and storage of specimens, 814 identification, 820 Granulicatella antimicrobial susceptibilities, 430, 1352 antimicrobial susceptibility testing, 1317, 1325–1326 clinical significance, 424 description of genus, 423 direct examination, 425 epidemiology and transmission, 423 identification, 426, 428, 429 interpretation of results, 431 isolation procedures, 425 taxonomy, 422–423 Granulicatella adiacens, 422, 430 antimicrobial susceptibility testing, 1325 blood culture, 18 endocarditis, 229 identification, 426 Granulicatella elegans, 422 antimicrobial susceptibility testing, 1325 identification, 426 Granuloma, tick, 2515 Granuloma inguinale, Klebsiella pneumoniae subsp. granulomatis and, 718 Granulomatous amebic encephalitis (GAE), Acanthamoeba and, 2391, 2394 Granulomatous hepatitis, Mycobacterium scrofulaceum and, 544 Graphium, 2154, 2159, 2180 Graves’ disease, Yersinia enterocolitica and, 742 Gravity displacement sterilizers, 204 Gray baby syndrome, 1193 Greasy pig syndrome, 360 Green bottle fly, 2517

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 85

GreeneChip, 144, 241 Gregg, McAlister, 1526 Grepafloxacin, 1179 Grimontia, 762, 763 Grimontia hollisae, 764, 766–769, 996 Griseofulvin, 2145, 2230 Grocott’s modification of the Gomori methenamine silver stain, 1958 Ground itch, 2456 Group A streptococci (GAS) clinical significance, 385–386 rectal swab screening for, 303 specimen collection, transport, and handling, 299–300 subtyping, 139 whole-genome sequencing, 245 Group B streptococci (GBS) clinical significance, 386 isolation procedures, 389–390 screening for, 296 Growth medium with 10% fetal bovine serum, 1430 Growth promoters use in food animals, 1214 Guanarito virus, 1669, 1671, 1674 Guaroa virus, 1645 Guillain-Barré syndrome (GBS), 1635 arboviruses, 1647 Campylobacter, 1000 Guinea worm, 2507; see also Dracunculus medinensis Gulf War Syndrome, Mycoplasma and, 1093 Gum Listeria medium (gum base-nalidixic acid medium), 337 Gumma, Treponema and, 1059, 1061 Gymnascella, 2062, 2069, 2075, 2076 Gymnascella dankaliensis, 2062 Gymnascella hyalinospora, 2062, 2066, 2070, 2118 Gymnophallidae (family), 2290 Gymnophalloidea (superfamily), 2290 Gymnothecium, 1941 Gynecologic infections, Clostridium sordellii and, 946 gyrA Aeromonas, 1326 Campylobacter, 1327 Mycobacterium, 581, 1361 Neisseria gonorrhoeae, 1323, 1383 Neisseria meningitidis, 1324 Vibrio cholerae, 1331 gyrB, Mycobacterium, 581, 1361 H. pylori Quick test, 1019 H antigen, Salmonella, 703–704 H broth, 337 H2S production, 615 HAART (highly active antiretroviral therapy), 1922, 2442 HACEK group, 676–677 antimicrobial susceptibilities, 661–662 antimicrobial susceptibility testing, 1317, 1328 β-lactamase testing, 1328 clinical significance, 654–655 identification, 653 Haemaphysalis, 2515 Anaplasma, 1138 Ehrlichia, 1138 key to identification, 2514 Haemaphysalis leachi, 2515 Haematobacter, 615, 623, 820 Haematobacter massiliensis, 632–633, 820 Haematobacter missouriensis, 632–633, 820 Haemophilus, 652, 667–680

n lxxxv

antimicrobial susceptibilities, 678–679, 1172–1174, 1176, 1181 resistance rates, 678–679 testing algorithm, 679 testing methods, 679, 1265 clinical significance, 669–670 collection, transport, and storage of specimens, 670–671 description of genus, 667–668 detection in blood, 20 differentiation of Francisella from, 852 direct examination, 671–672 antigen detection, 671 microscopy, 671 molecular techniques, 671–672 epidemiology and transmission, 668–669 evaluation, interpretation, and reporting of results, 679–680 identification, 675–677 commercial biochemical systems, 676 conventional biochemical tests, 676 mass spectrometry, 676–677 molecular identification, 677 problems with, 677 X and V factor growth requirements, 675–676 isolation procedures, 656, 672–675 colony appearance, 673–675 media, 672–674 serologic tests, 678 taxonomy, 667–668 typing systems, 677–678 capsular serotyping and biotyping, 677– 678 molecular methods, 678 Haemophilus aegyptius, 667–668, 670, 672, 674, 676–677, 680 Haemophilus aphrophilus, see Aggregatibacter aphrophilus Haemophilus ducreyi, 297, 667–668, 670–672, 674–675, 679–680, 1063–1064, 1183 Haemophilus haemolyticus, 667–668, 670, 674 Haemophilus influenzae, 667–680 antibiotic resistance, 1223, 1228, 1231, 1234, 1320–1321 antimicrobial susceptibilities, 678–679, 1174–1175, 1177–1178, 1180, 1182, 1184, 1187, 1190, 1192– 1193, 1195, 1197 BLNAR (β-lactamase negative and ampicillin resistant), 679 resistance rates, 678–679 testing algorithm, 679 testing methods, 679 antimicrobial susceptibility testing, 1320– 1322 commercial test methods, 1322 incidence of resistance, 1320–1321 reference test methods, 1321–1322 strategies for testing and reporting of results, 1322 β-lactamase, 1302, 1320–1322 β-lactamase tests, 1302 biotypes, 669 clinical significance, 669–670 collection, transport, and storage of specimens, 670–671 colony appearance, 673–674 description, 667–668 direct examination, 671–672 antigen detection, 671 microscopy, 671 molecular techniques, 671–672 epidemiology and transmission, 668–669

lxxxvi

n

SUBJECT INDEX

Haemophilus influenzae (continued) evaluation, interpretation, and reporting of results, 679–680 genome sequencing, 240–241 identification, 675–677 commercial biochemical systems, 676 conventional biochemical tests, 676 mass spectrometry, 676–677 molecular identification, 677 problems with, 677 X and V factor growth requirements, 675–676 in cystic fibrosis patients, 299 isolation procedures, 672–675 nontypeable (NTHi), 667–670, 678–679 reference strains, 1315 serologic tests, 678 taxonomy, 667–668 type b (Hib), 301, 668–669, 677 typing systems, 677–678 capsular serotyping and biotyping, 677– 678 molecular methods, 678 Wayson stain, 323 Haemophilus parahaemolyticus, 667–668, 674 Haemophilus parainfluenzae, 667–668, 670, 672, 674–678, 1172, 1321 Haemophilus paraphrohaemolyticus, 667–668, 674 Haemophilus paraphrophilus, see Aggregatibacter aphrophilus Haemophilus pittmaniae, 667–668 Haemophilus sputorum, 667–668 Haemophilus test medium, 337, 1269, 1321 Haemosporida (order), 2287, 2338 Hafnia epidemiology, transmission, and clinical significance, 721 taxonomy, 714 Hafnia alvei, 687, 693, 716, 721, 726 Hafnia paralvei, 721, 726 Hain MTBDRplus assay, 1368 Hain MTBDRsl assay, 1368 Hair specimen, for fungi, 1945, 1947, 1949 Halofantrine, 2538–2539 adverse effects, 2539 mechanism of action, 2539 pharmacokinetics, 2539 spectrum of activity, 2539 Halzoun, 2516 Hand disinfection, 186–187 Hand hygiene, 175, 186–187 Hand-foot-and-mouth disease, 1540–1541 Hanks’ balanced salt solution, 1422 Hansen’s disease, see Leprosy Hansenula angusta, 1993 Hansenula fabianii, 1993 Hantaan virus, 1660–1662 Hantavax, 1662 Hantavirus, 1660–1665 antigen detection, 1663 antiviral susceptibilities, 1665 biosafety, 1663 clinical significance, 1662 collection, transport, and storage of specimens, 1408, 1662–1663 description of agents, 1660–1661 detection and identification methods, 1433 direct examination, 1663 electron microscopy, 1660–1661, 1663 epidemiology and transmission, 1660–1661 evaluation, interpretation, and reporting of results, 1665

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 86

genomes, 1660 identification, 1664 genetic methods, 1664 serologic methods, 1664 isolation procedures, 1663–1664 laboratory criteria for diagnosis of hantavirus pulmonary syndrome, 1660 laboratory tests suggested for, 125 microscopy, 1663 nucleic acid detection, 1663 rodent-borne, 1665 serologic tests, 1664–1665 taxonomy, 1660 therapy, 1662 typing systems, 1664 vaccines, 1662 Hantavirus (genus), 1399, 1660, 1664 Hantavirus cardiopulmonary syndrome (HCPS), 1660 Hantavirus pulmonary syndrome (HPS), 1660–1665 Haplorchis, 2482, 2490 Harada-Mori filter paper strip culture, 2322– 2323, 2457 HardyCHROM Candida, 1960 HardyCHROM CRE agar, 337 HardyCHROM ESBL agar, 337 HardyCHROM MRSA, 1293 Harpellales (order), 2087 Hartland virus, 1645 Hartley’s digest broth, 337 Hartmannella, 2387 Hashimoto’s thyroiditis human herpesvirus 6 (HHV-6), 1756 Yersinia enterocolitica, 742 HAV, see Hepatitis A virus HAVpur, 1585 Havrix, 1584, 1590 HBoV, see Human bocavirus HBV, see Hepatitis B virus HBV trender, 1849 hc2 High-Risk HPV DNA test, 1414 HCoV-229E, 1565–1570, 1572–1578 HCoV-HKU1, 1565–1569, 1572–1578 HCoV-NL63, 1565–1578 HCoV-OC43, 1565–1569, 1572–1578 HCV, see Hepatitis C virus HCV DNA Chip v2.0, 1606 HDA (helicase-dependent amplification), 66–68 HDV, see Hepatitis D virus Head and neck tumors, HPV, 1786 Head louse, 2510–2511 Headache Anaplasma phagocyrophilum, 1139 arboviruses, 1647 arenaviruses, 1673–1674 Balantidium coli, 2417 centipede bites, 2520 Chlamydia psittaci, 1109 Corynebacterium diphtheriae, 480 Cryptosporidium, 2438 Cyclospora cayetanensis, 2428 Cystoisospora belli, 2428 Ehrlichia chaffeensis, 1138 filoviruses, 1674 fosfomycin, 1196 herpes simplex virus (HSV), 1689 Histoplasma capsulatum, 2114 Hymenolepis nana, 2476 influenza virus, 1471 linezolid, 1191 malaria, 2339

Mansonella, 2468 Orientia, 1124 Parastrongylus, 2498 phaeohyphomycoses, 2161 quinolones, 1180 Rickettsia, 1124 rifaximin, 1195 sulfonamides, 1192 Taenia solium, 2476 tetracyclines, 1187 Trichinella, 2495 variola virus, 1830 Health care-associated infections (HAIs), 106–116; see also Nosocomial infections Aspergillus, 2031–2033 common pathogens, 106, 108 costs, 107 definitions, 106, 107 emerging issues, 115–116 Enterobacteriaceae, 715 Enterococcus, 406–407 hospital infection prevention program, 107–110 antimicrobial stewardship, 110 infection prevention committee, 108– 109 process surveillance, 110 surveillance programs, 109–110 human metapneumovirus, 1509 infection rates, 106 infection sites, 106 morbidity and mortality, 106–107, 108 mucormycosis, 2089 Mycobacterium, 600 outbreaks, 126 Pantoea, 719 role of clinical microbiology laboratory in infection prevention, 110–116 identification and susceptibility testing of pathogens, 110–111 laboratory information system, 111 molecular typing, 112–113 organism banking and typing, 113 outbreak recognition and investigation, 112 rapid diagnostic testing, 111–112 reporting laboratory data, 112 specimen collection and transport, 110 surveillance cultures, 113–115 vancomycin-resistant enterococci (VRE), 406–407 HealthMap, 128 Hearing loss cytomegalovirus, 1719 rubella, 1526 Heart infusion agar, 337 Heartburn, Giardia duodenalis and, 2411 Heartland virus, 1653 Heartworm, see Dirofilaria immitis Heat map, 143 Hecht pneumonia, 1520 Hecolin, 1586, 1590 Hektoen enteric agar, 337 Helcobacillus chemotaxonomic features, 475 clinical significance, 479 description of genus, 477 identification, 438, 484, 494 Helcobacillus massiliensis, 494 Helcococcus clinical significance, 425 identification, 428, 429 isolation procedures, 425

SUBJECT INDEX Helcococcus kunzii, 425, 427, 428 Helcococcus pyogenes, 425, 428 Helcococcus sueciensis, 425, 428 Helicase-dependent amplification (HDA), 66–68 Helicobacter, 1013–1024 antimicrobial susceptibility, 1022–1023 clinical significance, 1017–1018 enterohepatic helicobacters, 1017–1018 H. pylori, 1017 HHLO, 1017 collection, transport, and storage of specimens, 1018 blood specimens, 1018 fecal specimens, 1018 gastric biopsy specimens, 1018 saliva, 1018 urine, 1018 description of agents, 1013–1014, 1016 direct examination, 1018–1020 H. pylori fecal antigen detection, 1019 microscopy, 1018–1019 nucleic acid detection, 1019–1020 urea breath test (UBT), 1019 urease testing of biopsy specimens, 1019 enterohepatic helicobacters antimicrobial susceptibility testing, 1023 characteristics of, 1016 clinical significance, 1017–1018 epidemiology and transmission, 1017 identification, 1021 isolation procedures, 1020 nucleic acid detection, 1020 phylogenetic tree, 1015 epidemiology and transmission, 1014– 1015, 1017 enterohepatic helicobacters, 1017 H. pylori, 1015, 1017 HHLO, 1017 evaluation, interpretation, and reporting of results, 1023–1024 HHLO (Helicobacter heilmannii-like organisms) antimicrobial susceptibility testing, 1023 clinical significance, 1017 epidemiology and transmission, 1017 isolation procedures, 1020 microscopy, 1018 nucleic acid detection, 1020 taxonomy, 1013 identification, 994–997, 1020–1021 isolation procedures, 1020 phylogenetic tree, 1015 serologic tests, 1021–1022 detection of H. pylori antibody in blood, 1021–1022 detection of H. pylori antibody in urine and saliva, 1022 taxonomy, 1013 typing systems, 1021 Helicobacter acinonychis, 1013–1016 Helicobacter anseris, 1014–1016 Helicobacter aurati, 1014–1016 Helicobacter baculiformis, 1014–1016 Helicobacter bilis, 996, 1013–1017, 1020– 1021 Helicobacter bizzozeronii, 996, 1013–1016, 1020, 1023 Helicobacter bovis, 1014–1017 Helicobacter brantae, 1014–1016 Helicobacter canadensis, 996, 1013–1018, 1020–1021 Helicobacter canis, 996, 1013–1017, 1020, 1023–1024

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 87

Helicobacter cetorum, 1014–1016 Helicobacter cholecystus, 1014–1016 Helicobacter cinaedi, 994, 996–997, 1013– 1017, 1019–1024 Helicobacter cynogastricus, 1014–1016 Helicobacter equorum, 1014–1016 Helicobacter felis, 1013–1017 Helicobacter fennelliae, 994, 996–997, 1013– 1018, 1020–1024 Helicobacter ganmani, 1014–1016 Helicobacter heilmannii, 996, 1013–1017, 1020, 1023 Helicobacter hepaticus, 1014–1016, 1018 Helicobacter macacae, 1014–1016 Helicobacter marmotae, 1014–1016 Helicobacter mastomyrinus, 1014–1016 Helicobacter mesocricetorum, 1014–1016 Helicobacter muridarum, 1014–1016 Helicobacter mustelae, 1013–1016 Helicobacter nemestrinae, 1014–1016 Helicobacter pametensis, 1014–1016 Helicobacter pullorum, 996, 1005, 1013–1018, 1020–1021, 1023–1024 Helicobacter pylori, 994, 996, 1013–1024 antibiotic resistance, 1231, 1328–1329 antibiotic therapy and relevance of resistance, 1022 antimicrobial susceptibility, 1022–1023, 1180, 1183, 1195 genotypic susceptibility testing in feces, 1023 genotypic susceptibility testing of cultures and biopsy specimens, 1023 phenotypic susceptibility testing of cultures, 1022–1023 antimicrobial susceptibility testing, 1317, 1328–1329 commercial test method, 1329 incidence of resistance, 1328–1329 reference test method, 1329 strategies for testing and reporting of results, 1329 characteristics of, 1016 clinical significance, 1017 collection, transport, and storage of specimens, 303, 1018 blood specimens, 1018 fecal specimens, 1018 gastric biopsy specimens, 1018 saliva, 1018 urine, 1018 description, 1013–1014 direct examination, 1018–1020 H. pylori fecal antigen detection, 1019 microscopy, 1018–1019 nucleic acid detection, 1019–1020 urea breath test (UBT), 1019 urease testing of biopsy specimens, 1019 epidemiology and transmission, 1014– 1015, 1015, 1017 evaluation, interpretation, and reporting of results, 1023–1024 identification, 1020–1021 in gastric microbiome, 229–230 isolation procedures, 1020 nucleic acid detection, 1019–1020 in feces, 1019 in gastric biopsy specimens, 1019 phylogenetic tree, 1015 serologic tests, 1021–1022 detection of antibody in blood, 1021– 1022 detection of antibody in urine and saliva, 1022

n lxxxvii

taxonomy, 1013 typing systems, 1021 Helicobacter rodentium, 1014–1016 Helicobacter salomonis, 1013–1017 Helicobacter suis, 996, 1014–1018, 1020 Helicobacter trogontum, 1013–1016 Helicobacter typhlonius, 1014–1016 Helicobacter winghamensis, 1014–1018, 1020– 1021 Helicobacteraceae (family), 1013 Helminths anthelminthic agents, 2529–2536 commercial kits for immunodetection of serum antibodies, 2296 less common, 2493–2503 cestodes, 2499–2503 collection, transport, and storage of specimens, 2493 nematodes, 2493–2501 recovery and identification techniques, 2323 taxonomy and classification, 2288–2291 acanthocephalans, 2291 cestodes, 2288, 2291 nematodes, 2288–2289 trematodes, 2288, 2290 Helcococcus, 422–423 Helcococcus kunzii, 423 Helcococcus ovis, 423 Helcococcus pyogenes, 423 Helcococcus sueciensis, 423 Hemadsorption (HAD) test mumps virus, 1494 parainfluenza virus, 1491 poxviruses, 1833 test procedure, 1423 Hemagglutination assays, 96, 1832–1833 parvovirus B19, 1821 Hemagglutination inhibition (HI), 96 adenovirus, 1776–1777 arboviruses, 1653 influenza viruses, 1480–1481 mumps virus, 1494–1495 parainfluenza virus, 1492 polyomavirus, 1810 poxviruses, 1835 rubella virus, 1529 Hemagglutinin (HA), influenza virus, 1470– 1471, 1476, 1479–1482 Hematologic disease specimens, 1406 Hematopoietic stem cell transplantation (HSCT) adenoviruses, 1771–1772, 1775, 1777 Fusarium, 2065, 2067 human herpesvirus 6 (HHV-6), 1756 respiratory syncytial virus and, 1501 Hematoxylin and eosin (H&E) stain, for fungi, 1965, 1970, 1974, 1975, 1976 Hematoxylin stains, for parasitology, 2314, 2316 Hematuria, BK polyomavirus and, 1804 Hemimetabolous development, 2508, 2510 Hemiplegia, human herpesvirus 7 (HHV-7) and, 1761 Hemiptera (order), 2507–2509, 2522 Hemolytic anemia cephalosporins, 1175 Mycoplasma, 1091 nitrofurantoin, 1196 penicillins, 1173 rifampin, 1195 sulfonamides, 1192 viruses, specimens and methods for detection of, 1406

lxxxviii n

SUBJECT INDEX

Hemolytic-uremic syndrome (HUS), 688, 692, 696–697 Hemophagocytic lymphohistiocytosis syndrome, 1138 Hemoptysis, dirofilariasis and, 2499 Hemorrhage arboviruses, 1647 arenaviruses, 1673 filoviruses, 1674 Hemorrhage-hepatitis syndrome, enterovirus, 1540 Hemorrhagic colitis, Klebsiella oxytoca and, 718 Hemorrhagic cystitis adenovirus, 1778 adenoviruses, 1772, 1773 BK polyomavirus, 1804–1805, 1811 enteric adenoviruses, 1622 viruses, specimens and methods for detection of, 1408 Hemorrhagic fever viruses (HFVs), 125, 222 Hemorrhagic fever with renal syndrome (HFRS), 1660–1665 Hemorrhagic syndrome, Lonomia achelous and, 2518 Hemozoin, 2347 Hendersonula, 2146 Hendersonula toruloidea, 2153 Henipavirus (genus), 1398 Hepacivirus (genus), 1399, 1599 Hepadnaviridae (family), 1398, 1400–1401, 1841 Hepatic abscess Ascaris lumbricoides, 2451 Pediococcus, 424 Hepatitis adenoviruses, 1772, 1773 arboviruses, 1647 Bartonella, 876 Campylobacter, 1000 cytomegalovirus, 1719 enteric adenoviruses, 1622 epidemic, 1841 Epstein-Barr virus, 1739 hepatitis A virus, 1589–1591 hepatitis B virus, 1844–1845, 1852–1853 hepatitis C virus, 1599–1601 hepatitis D virus, 1854–1855 hepatitis E virus, 1589–1591 herpes simplex virus (HSV), 1689 Histoplasma capsulatum, 2114 human herpesvirus 6 (HHV-6), 1755– 1756 liver trematodes, 2489 macrolides, 1183 Mansonella, 2468 measles, 1521 microsporidia, 2210, 2213 nitrofurantoin, 1196 penicillins, 1173 rifampin, 1195 varicella-zoster virus, 1705 viruses, specimens and methods for detection of, 1406 Hepatitis A infection, 1589–1591 Hepatitis A virus (HAV), 1584–1594 antigen detection, 1591 clinical significance, 1589–1591 clinical presentation and course, 1589– 1590 dose response to infection, 1590 vaccines and antiviral agents, 1590– 1591 description of agent, 1584–1585

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 88

detection and identification methods, 1433 direct examination, 1591 electron microscopy, 1591 epidemiology and transmission, 1586–1588 export from hepatocytes, 1586 genome replication and proteins, 1585 identification and typing systems, 1592 isolation procedures, 1592 laboratory tests suggested for, 125 nucleic acid detection, 1591 serologic tests, 1592 specimen collection and handling, 1406, 1413, 1591 taxonomy, 1584 vaccine, 1584–1585, 1589–1590 Hepatitis B infection acute, 1843–1844, 1852 chronic, 1843–1845, 1852–1853, 1880 epidemiology and transmission, 1843–1844 laboratory techniques and control interventions used in significant outbreaks, 126 laboratory tests suggested for, 125 laboratory-acquired infections, 177 worldwide distribution of chronic, 1844 Hepatitis B virus (HBV) antigen detection, 1845–1848 antiviral agents, 1880–1882 nucleoside/nucleotide analogues, 1880– 1882 antiviral resistance, 1917, 1923–1924 antiviral resistance mechanisms, 1899– 1900 antiviral susceptibility testing, 1851–1852, 1920–1921 carriers, inactive, 1853 clinical significance, 1844–1845 collection, transport, and storage of specimens, 1406–1407, 1412, 1845 description of agent, 1841–1843 detection and identification methods, 1433 direct detection, 1845–1849 epidemiology and transmission, 1843–1844 evaluation, interpretation, and reporting of results, 1852–1854 genome, 1841–1842 genotypes, 1849–1850, 1900 HBcAg (hepatitis B core protein antigen), 1841–1843, 1845, 1848–1850 HBeAg (hepatitis B e antigen), 1841– 1843, 1845, 1851–1853 HBsAg (hepatitis B surface antigen), 1841–1847, 1850–1853 HDV coinfection, 1854–1855 history, 1841 identification, 1849 isolation, 1849 markers, 1843, 1852 microscopy, 1845 nucleic acid detection, 1411, 1846, 1849 occupational exposure, 1845 serologic tests, 1847–1851 anti-HBe, 1851 anti-HBs, 1850–1851 commercial systems, 1847–1848 IgM anti-HBc, 1849–1850 total anti-HBc, 1850 taxonomy, 1841 typing systems, 1849–1850 vaccine, 1585, 1843, 1845, 1849–1851, 1853 virion morphology, 1841–1842

Hepatitis C infection, 1599–1601 acute clinical features of, 1599–1600 diagnosis, 1603 interpretation of results in, 1609–1610 antiviral resistance, 1901–1903, 1917, 1923–1924 antiviral susceptibility, 1609 chronic clinical features, 1600–1601 diagnosis, 1603 interpretation of results in, 1610–1611 screening recommendations, 1607 diagnosis, 1602–1609 epidemiology and transmission, 1599 laboratory-acquired infections, 177 treatment, 1601–1602, 1901 direct-acting antiviral drugs (DAAs), 1601–1602, 1610–1611 HCV RNA quantification to define therapeutic response, 1610–1611 nucleic acid tests (NATs) in management of chronic hepatitis C therapy, 1603–1605 viral load, 1601–1602, 1610 Hepatitis C virus (HCV), 1599–1611 antibody testing, 1409 antigen detection, 1603 antiviral agents, 1878–1880 combination therapies, 1880 interferon, 1878–1879 polymerase inhibitors, 1879–1880 protease inhibitors, 1879 ribavirin, 1878–1879 table of agents, 1879 antiviral resistance, 1917, 1923–1924 antiviral resistance mechanisms, 1900– 1903 interferon resistance, 1901–1902 nonnucleoside inhibitor (NNI) resistance, 1902–1903 NS5A inhibitor resistance, 1902–1903 nucleoside inhibitor resistance, 1902– 1903 protease inhibitor resistance, 1901–1902 ribavirin resistance, 1901–1902 antiviral susceptibility, 1609 antiviral susceptibility testing, 1921 clinical significance, 1599–1601 collection, transport, and storage of specimens, 1406, 1411–1412, 1414, 1602 description, 1599 detection and identification methods, 1433 direct examination, 1602–1605 discovery of, 240 endoscope contamination outbreak, 199 epidemiology and transmission, 1599 evaluation, interpretation, and reporting of results, 1609–1611 in acute hepatitis C, 1609–1610 in chronic hepatitis C, 1610–1611 genome and protein coding scheme, 1599–1600 genotypes, 1599, 1605–1606, 1610, 1901 genotyping, 1605–1607 identification, 1605 isolation procedures, 1605 microscopy, 1602–1603 nucleic acid tests (NATs), 1411, 1603– 1605 commercial HCV RNA qualitative tests, 1603–1604

SUBJECT INDEX commercial HCV RNA quantitative tests, 1604–1605 diagnosis of acute infection, 1603 diagnosis of chronic infection, 1603 management of chronic hepatitis C therapy, 1603–1605 nucleic acid preparation for, 1604 serologic tests, 1606–1609 taxonomy, 1599 testing algorithm, 1609 therapy, 1601–1602, 1901 molecular methods of monitoring response, 76–77 Hepatitis D infection, 1854–1855 Hepatitis D virus (HDV), 1854–1855 clinical significance, 1854–1855 collection, transport, and storage of specimens, 1406, 1855 description of agent, 1854 detection and identification methods, 1433 direct detection, 1855 epidemiology and transmission, 1854 evaluation, interpretation, and reporting of results, 1855 HBV coinfection, 1854–1855 serologic tests, 1855 taxonomy, 1854 typing, 1855 Hepatitis E infection, 1589–1591 Hepatitis E virus (HEV), 1584–1594 antigen detection, 1591 clinical significance, 1589–1591 clinical presentation and course, 1589– 1590 dose response to infection, 1590 vaccines and antiviral agents, 1590– 1591 collection, transport, and storage of specimens, 1406, 1591 description of agent, 1585–1586 detection and identification methods, 1433 direct examination, 1591–1592 electron microscopy, 1591 epidemiology and transmission, 1586–1589 genome replication and proteins, 1585 genotypes, 1586 identification and typing systems, 1592 isolation procedures, 1592 laboratory tests suggested for, 125 nucleic acid detection, 1591–1592 serologic tests, 1592–1594 taxonomy, 1584 vaccine (candidate), 1586, 1590 Hepatobiliary disease, Helicobacter and, 1018 Hepatocellular carcinoma (HCC), hepatitis B virus and, 1844–1845, 1852 Hepatomegaly Leishmania, 2359 Trypanosoma brucei, 2366 Trypanosoma cruzi, 2362 Trypanosoma lewisi, 2368 Hepatosplenic schistosomiasis, 2479, 2484 Hepatosplenomegaly, HHV-6, 1755 Hepatotoxicity nitrofurantoin, 1196 telithromycin, 1185 Hepatovirus (genus), 1399, 1551, 1584 HEPES, 1423 Hepeviridae (family), 1399–1401, 1584 Hepevirus (genus), 1399, 1584 Herbaspirillum, 615, 630–631, 994, 997 Herpangina, enterovirus, 1540–1541

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 89

Herpes B virus, 1687, 1696–1697 clinical significance, 1697 collection, transport, and storage of specimens, 1697 description of agent, 1696–1697 identification, 1697 reference laboratories, 1697 serodiagnosis, 1697 Herpes simplex virus (HSV), 1687–1696 antigen detection, 1691–1692 antiviral resistance, 1894–1895, 1917 antiviral susceptibilities, 1695 antiviral susceptibility testing, 1916, 1919 clinical significance, 1688–1689 antiviral therapy, 1689 asymptomatic or subclinical infection, 1688 central nervous system disease in immunocompetent host, 1689 immunocompromised host, 1689 latency and recurrent disease, 1688 neonatal herpes, 1688–1689 ocular infection, 1689 primary infection, 1688 systemic disease in hospitalized adults, 1689 collection, storage, and transport of specimens, 1689–1690 collection and handling, 1406–1408, 1415 storage and processing, 1412–1413 cytopathic effect (CPE), 1690, 1692 description of agent, 1687–1688 detection, 1427 detection and identification methods, 1434 detection tests, interpretation of, 1695– 1696 DFA and IFA reagents for the detection of, 1425 diagnostic tests, 1693 direct detection, 1690–1692 epidemiology and transmission, 1688 evaluation, interpretation, and reporting of results, 1695–1696 identification and typing, 1692–1693 immunofluorescence in H&V-Mix cells, 1429 isolation procedures, 1692 microscopy, 1691 N-Docosanol, 1689 nucleic acid tests, 1690–1691 rapid cell culture, 1426 serologic tests, 1693–1694 taxonomy, 1687 thymidine kinase, 1894–1895 TORCH (toxoplasmosis, other, rubella, cytomegalovirus, and herpes simplex virus) panels, 1530 transport medium for, 1409 type-specific serology, interpretation of, 1696 virion morphology, 1687 Herpes virus LC-PCR kits, 1707 Herpes zoster, 1705–1706, 1709, 1712; see also Varicella-zoster virus postherpetic neuralgia, 1705–1706 visceral zoster, 1705–1706, 1709 zoster sine herpete, 1705–1706, 1709 HerpeSelect EIA, 1694 HerpeSelect ELISA, 1693–1694 HerpeSelect Express IgG, 1693 HerpeSelect Immunoblot, 1693 Herpesvirales (order), 1398, 1402

n lxxxix

Herpesviridae (family), 1738 cytomegalovirus (CMV), 1718–1732 herpes simplex virus (HSV), 1687–1696 immunofluorescence detection, 1429 taxonomic classification, 1398, 1400 varicella-zoster virus, 1704–1713 virion morphology, 1401 Herpesvirus simiae, see Herpes B virus Herpesviruses antiviral agents, 1882–1886 antiviral resistance, 1917–1919 antiviral resistance mechanisms, 1894– 1896 Herpetic lesion specimens, 1408 Heteroduplex mobility analysis (HMA) hepatitis C virus, 1607 respiratory syncytial virus, 1505 Heterokonta (phylum), 2203 Heterolobosea (class), 2287, 2387 Heterophyes, 2320, 2482 Heterophyes heterophyes, 2449, 2484, 2490 Heterophyiasis, 2482 Heterophyidae (family), 2290, 2482, 2490 Heterothallic, 1936, 1937, 1941 Heterotrophs, 1935 HEV, see Hepatitis E virus HEV IgM ELISA 3.0, 1592–1593 HEV IgM RPOC test, 1593–1594 Hexadecyloxypropyl cidofovir, 1777 HGA (human granulocytic anaplasmosis), 1138–1139, 1142–1145 Hg-PVA, 2311–2312 HI (human intestinal spirochetosis), 1055, 1058–1059, 1061–1062 HiCrome aureus agar base with egg yolk tellurite (Staphylococcus aureus agar, HiCrome), 337 HiCrome Candida differential agar, 1960, 1999 HiCrome Listeria agar base, modified, with moxalactam (Listeria HiCrome agar base, modified), 337–338 HiCrome MeReSa agar with methicillin, 338 HiCrome RajHans medium, 338 HiCrome RajHans medium, modified, 338 HiCrome Salmonella agar, 338 HiCrome UTI agar, 338 HiFluoro Pseudomonas agar base, 338 High Pure/Cobas TaqMan v2.0, 1604–1605 Highly active antiretroviral therapy (HAART), 1922, 2442 High-performance liquid chromatography (HPLC), 1380 High-resolution computerized tomography (HRCT), Pneumocystis and, 2019 Hill, Bradford, 246 Hilum, 1941 Himasthla, 2482 Hippurate hydrolysis test, 318, 392 Histamine, 2518 Histidine-rich protein 2 (HRP-2) test, for parasites, 2336 Histomonas, 2412 Histopathology cytomegalovirus (CMV), 1722 mucormycosis, 2090 Histoplasma, 1937, 1939, 1948, 2109 Histoplasma capsulatum, 1935, 1938, 1939, 2109–2123 African histoplasmosis, 2114 antibody detection, 1971 antifungal susceptibilities, 2121–2122, 2224 antigen detection, 1977–1978, 2116

xc

n

SUBJECT INDEX

Histoplasma capsulatum (continued) biosafety, 2117 blood culture, 18 blood specimens, 2307 clinical significance, 2114 culture for mold phase, 2117 culture for yeast phase, 2118 description of agents, 2110–2111 detection in blood, 21, 2336 direct examination, 2115–2117 endophthalmitis, 1949 epidemiology and transmission, 2113 evaluation, interpretation, and reporting of results, 2121–2123 identification, 2118 isolation, 2117–2118 media, 1959 microscopy, 1966, 1976, 2110–2111, 2115–2116 nucleic acid detection, 2117 serologic tests, 2120 specimen collection, transport, and processing, 1948–1949, 1951, 2115 staining, 1957 taxonomy, 2109 typing systems, 2119 Histoplasma capsulatum var. capsulatum, 2109 Histoplasma capsulatum var. duboisii, 2109 Histoplasma capsulatum var. farciminosum, 2109 Histoplasma polysaccharide antigen (HPA), 2116, 2121 Histoplasmin, 1971 Histoplasmosis, 1971, 1977, 2109–2123 African, 2114 antigen detection, 2116 clinical significance, 2114 description of agents, 2110–2111 epidemiology and transmission, 2113 evaluation, interpretation, and reporting of results, 2121–2123 nucleic acid detection, 2117 serologic tests, 2120 specimens for, 1947, 1949–1950, 2115 Histotoxic skin and soft tissue infections clostridial, 944–946 HIV, see Human immunodeficiency virus (HIV) HIV RNA assay, 1409 HME (human monocytic ehrlichiosis), 1138–1142, 1144–1145 HMP Reference Genomes Catalog, 233 HMPV, see Human metapneumovirus Hodgkin’s lymphoma, Epstein-Barr virus and, 1738, 1740–1741 Holdemania, 921 Holdemania filiformis, 922, 927, 930 Holoblastic, 1940–1941, 1941 Holoblastic conidiogenesis, 2058 Holocyclotoxin, 2516 Hologic/Gen-Probe assay, for Trichomonas vaginalis, 2415 Holometabolous development, 2505 Holothyrida (order), 2511 Holozoa (division), 1936 Homothallic, 1936, 1941 Homothallic ascomycetes, 2075 clinical significance, 2075 description, 2069 key phenotypic features, 2062 Honeybees, 2518 Hook effect, 99–100 Hooke, Robert, 5 Hookworm, 2454–2456

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 90

clinical significance, 2456 description, 2454 eggs, 2449, 2452, 2454 larvae, 2452, 2454 worms, 2454 detection, 2320, 2323, 2329, 2331 diagnosis, 2456 epidemiology and prevention, 2454 taxonomy, 2454 transmission and life cycle, 2454, 2456 treatment, 2455–2456, 2531–2532 Horie arabinose ethyl violet broth, 338 Hormographiella, 2062, 2071 Hormographiella aspergillata, 2062, 2075 Hormographiella verticillata, 2062 Hormonema, 2153, 2159 Hormonema dematioides, 2155, 2158, 2163 Hornets, 2518, 2522 Horse/deerflies, 2505 Hortaea, 2155–2156 Hortaea werneckii, 2144, 2147–2148, 2154, 2156, 2161–2162 Hospital Infection Control Practices Advisory Committee, 406 Hospitalized adults, herpes simplex virus (HSV) in, 1689 HotSHOT DNA extraction, 2523 House mouse mite, 2511 Housefly, 2513, 2517 Hoverfly, 2517 Hoyle medium, 338 HPeV, see Parechoviruses HPLC identification, of Mycobacterium, 601 HPV, see Human papillomavirus HPV direct-flow chip, 1791 HPV infection in men (HIM), 1785 HRCT (high-resolution computerized tomography), Pneumocystis and, 2019 hSLAM molecule, 1522–1523 hsp65 gene, Mycobacterium, 580, 602–603 HSV, see Herpes simplex virus HSV1 HSV2 VZV R-gene, 1707 HT-2 toxin, 2190 HTLV-1/2 ChLIA, 1463 HTLV-1/2 EIA, 1463 HTLV-1-associated myelopathy/tropical spastic paraparesis (HAM/TSP), 1460–1462 HTLVs, see Human T-cell lymphotropic viruses HU (hemolytic-uremic syndrome), 688, 692, 696–697 Hülle cell, 1941 Human bocavirus (HBoV), 242, 1406–1407, 1433, 1618, 1823–1824 antigen detection, 1823 clinical significance, 1823 description, 1823 direct examination, 1823–1824 epidemiology and transmission, 1823 evaluation, interpretation, and reporting of results, 1824 isolation, 1824 microscopy, 1823 nucleic acid detection, 1824 serologic tests, 1824 taxonomy, 1818 typing, 1824 Human botfly, 2517 Human coronaviruses (HCoVs), 1407–1408, 1565–1578; see also Coronaviruses Human cytomegalovirus, see Cytomegalovirus

Human enterovirus, rhinoviruses distinguished from, 1551, 1554–1555, 1557–1559 Human enterovirus C, 1395–1396 Human granulocytic anaplasmosis (HGA), 1138–1139, 1142–1145 Human granulocytic ehrlichiosis (HGE), 2507, 2521, 2523 Human herpesvirus 1, see Herpes simplex virus Human herpesvirus 2, see Herpes simplex virus Human herpesvirus 3 (HHV-3), 1704; see also Varicella-zoster virus Human herpesvirus 4 (HHV-4), see EpsteinBarr virus Human herpesvirus 5 (HHV-5), 1718; see also Cytomegalovirus Human herpesvirus 6 (HHV-6), 1754–1761 antiviral susceptibilities, 1760 clinical significance, 1755–1756 brain infections, 1756 HIV-infected patients, 1756 primary infection, 1755–1756 therapy, 1756 transplant recipients, 1756 commercial testing, 1760 description of agent, 1754 detection and identification methods, 1434 diagnostic methods, 1757 direct examination, 1756–1759 epidemiology, 1754 evaluation, interpretation, and reporting of results, 1760–1761 future directions, 1764–1765 identification, 1759 immunocompromised patients, 1754–1756, 1760 isolation, 1757, 1759 latency, persistence, and transmission, 1754–1755 PCR, 1756–1757, 1759–1760 serologic tests, 1757, 1759–1760 specimen collection and handling, 1406– 1407, 1756 taxonomy, 1754 tissue distribution, 1754 typing systems, 1759 Human herpesvirus 7 (HHV-7), 1761–1762 antiviral susceptibilities, 1762 clinical significance, 1755, 1761 immunocompromised patients, 1761 primary infection, 1761 therapy, 1761 collection, transport, and storage of specimens, 1761 commercial testing, 1760 description of the agent, 1761 detection and identification methods, 1434 diagnostic methods, 1757–1758 direct examination, 1761 epidemiology, 1761 evaluation, interpretation, and reporting of results, 1762 future directions, 1764–1765 identification, 1762 isolation, 1757, 1761–1762 PCR, 1758–1760 serology, 1757, 1759–1760, 1762 taxonomy, 1761 tissue distribution, 1761 transmission, 1761

SUBJECT INDEX Human herpesvirus 8 (HHV-8), 1762–1765 antiviral susceptibilities, 1764 clinical significance, 1755, 1762–1763 Kaposi’s sarcoma, 1762–1763 multicentric Castleman’s disease, 1763 PEL, 1763 primary infection, 1762 therapy, 1763 transplant recipients, 1763 commercial testing, 1760 description of agent, 1762 detection and identification methods, 1434 diagnostic methods, 1758 direct detection, 1763 discovery of, 240 epidemiology, 1762 evaluation, interpretation, and reporting of results, 1764 future directions, 1764–1765 identification, 1764 isolation, 1764 latency and persistence, 1762 PCR, 1758–1760 serology, 1758–1760, 1764 specimen collection and handling, 1406, 1763 taxonomy, 1762 tissue distribution, 1762 transmission, 1762 typing, 1764 Human immunodeficiency virus (HIV), 1436–1451 antiviral agents, 1869–1878 entry inhibitors, 1871, 1877 integrase strand transfer inhibitors, 1872, 1877–1878 nonnucleoside reverse transcriptase inhibitors (NnRTIs), 1870, 1873–1874 nucleoside/nucleotide reverse transcriptase inhibitors (NRTIs/ NtRTIs), 1869–1870, 1872–1873 protease inhibitors, 1871, 1874–1875 table of agents, 1870–1872 antiviral resistance, 1919–1920, 1923– 1924 antiviral resistance mechanisms CCR5 inhibitor resistance, 1897, 1899 fusion inhibitor resistance, 1897, 1899 integrase inhibitor resistance, 1897– 1899 intersubtype variation, 1899 non-nucleoside reverse transcriptase inhibitor (NNRTI) resistance, 1897–1898 nucleoside/nucleotide reverse transcriptase inhibitor (NRTI) resistance, 1896–1898 protease inhibitor resistance, 1897–1898 antiviral susceptibilities, 1447–1448 interpretation of resistance assays, 1450–1451 antiviral susceptibility testing, 1919–1920 clinical significance, 1438–1440 acute retroviral syndrome, 1438–1439 clinical latency, 1439 conventional immunoassays, 1446 disease progression to AIDS, 1439 initial screening tests, 1444–1446 therapy and vaccination, 1440 virological parameters during the course of HIV infection, 1438–1439 coinfection

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 91

Acanthamoeba, 2391 adenoviruses, 1771 Blastocystis hominis, 2406 Candida, 1993 Cryptococcus, 1993 Cryptosporidium, 2437–2438, 2442 Cystoisospora belli, 2428, 2430–2431 cytomegalovirus, 1719–1720 Dientamoeba fragilis, 2413 Entamoeba histolytica, 2403 Epstein-Barr virus, 1740 human herpesvirus 6 (HHV-6), 1756, 1760 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1762– 1764 human papillomavirus, 1784 human T-cell lymphotropic viruses (HTLVs), 1461 Leishmania, 2358–2360, 2362 microsporidia, 2212–2213, 2215–2216 mucormycosis, 2088 Mycoplasma genitalium, 1093 Pneumocystis, 2017–2020 polyomaviruses, 1804–1805, 1811–1812 syphilis, 1058–1059, 1073, 1075 Talaromyces marneffei, 2046 Toxoplasma gondii, 2381–2382 Trichomonas vaginalis, 2414–2415 Trypanosoma cruzi, 2363 collection, storage, and transport of specimens, 1440–1441 collection and handling, 1406–1407, 1412–1414 storage and processing, 1411 description of the agents, 1436–1437 detection and identification methods, 1434 diagnostic human immunodeficiency virus testing algorithm, 1449 direct detection, 1441–1443 epidemiology and transmission, 1438, 2505 evaluation, interpretation, and reporting of results, 1449–1451 use and interpretation of qualitative HIV RNA and DNA assays, 1449–1450 use and interpretation of resistance assays, 1450–1451 use and interpretation of serologic tests, 1449 use and interpretation of viral load assays, 1450 genotyping assays, 1447–1448 results, 1450–1451 historical perspective and origin, 1436 HIV classification, 1436 HIV RNA and DNA qualitative assays, 1441–1442 isolation procedures, 1443 laboratory-acquired infections, 177 nucleic acid tests, 1411 p24 antigen assays, 1441 phenotyping assays, 1448 results, 1451 point-of-care human immunodeficiency virus tests, 1445 serologic diagnosis of, 1408–1409 serologic tests, 1443–1447 alternative specimens for antibody testing, 1445–1446 rapid immunoassays, 1446–1447

n xci

screening for atypical and human immunodeficiency virus type 2 infections, 1446 supplemental assays for human immunodeficiency virus, 1446– 1447 structure and genomic organization, 1436– 1437 taxonomy, 1436 time course of appearance of laboratory markers for HIV-1 infection, 1439 tropism assays, 1448 viral load assays, 75, 77, 1442–1443 interpretation of, 1450 virion morphology, 1436–1437 Human influenza virus real-time RT-PCR detection and characterization panel, 1477 Human intestinal spirochetosis (HIS), 1055, 1058–1059, 1061–1062 Human metapneumovirus (HMPV), 242, 1508–1512 antigen detection, 1510–1511 antiviral susceptibilities, 1512 clinical significance, 1509 collection, transport, and storage of specimens, 1407–1408, 1509 cytopathic effect (CPE), 1511–1512 description of agent, 1508 detection and identification methods, 1434 DFA and IFA reagents for the detection of, 1425 direct examination, 1510–1511 direct fluorescent antibody (DFA) test, 1503 epidemiology and transmission, 1509 evaluation, interpretation, and reporting of results, 1512 identification, 1511 isolation procedures, 1511 microscopy, 1510 nucleic acid amplification tests (NAATs), 1506–1507 nucleic acid detection, 1511 rapid cell culture, 1426 seasonal distribution, 1510 serologic tests, 1511–1512 taxonomy, 1508 typing systems, 1511 Human microbiome, 226–233 colonic, 230–231 esophageal, 229 gastric, 229–230 gastrointestinal, 229–231 Human Microbiome Project, 226 oral, 227–229 phyla predominant by body site, 228 reference strain genomes, 232–233 respiratory tract, 231 skin, 231–232 small intestinal, 230 techniques for study of, 226–227 terminology, 227 vaginal, 231 Human Microbiome Project (HMP), 226, 246 Human monocytic ehrlichiosis (HME), 1138–1142, 1144–1145, 2507 Human papillomavirus (HPV), 1783–1796 alphapapillomavirus genotype diversity and clinical manifestations, 1784 antigen detection, 1788 clinical significance, 1785–1786

xcii

n

SUBJECT INDEX

Human papillomavirus (HPV) (continued) clinical manifestations of female anogenital infection, 1785 clinical manifestations of male and female oral infection, 1786 clinical manifestations of male anogenital infection, 1785 primary prevention, 1786 collection, transport, and storage of specimens, 1786–1788 collection and handling, 1406, 1408 storage and processing, 1412–1414 description of the agent, 1783 detection and identification methods, 1434 diagnostic applications, 1788 direct detection, 1788–1795 epidemiology and transmission, 1783–1785 evaluation, interpretation, and reporting of results, 1795–1796 future of testing, 1795–1796 genotyping systems FDA-approved, 1794 research-use-only assays, 1795 in situ hybridization tests for HPV detection, 1789 line probe for identification of 37 HPV genotypes, 69 microscopy, 1788 nucleic acid isolation and purification, 1789, 1792 exfoliated cells, 1789 formalin-fixed paraffin-embedded tissue, 1789 fresh frozen tissue, 1789 nucleic acid tests, 1788–1795 Aptima HPV assay, 1793 Cervista HR-HPV test, 1792–1793 Cobas HPV test, 1793 comparison of HPV assays for detection of CIN2+ and CIN3+, 1793– 1794 genotyping assays, 1794–1795 hc2 tests, 1792 methylation as biomarker, 1795 screening applications, 1786–1787 self-collection for HPV infection, 1788 serologic tests, 1795 subtyping by molecular methods, 76 taxonomy, 1783 vaccine, 1786 Human parainfluenza viruses (HPIVs), see Parainfluenza virus Human parvovirus 4 (PARV4), 1818, 1824 Human polyomavirus 6 (HPyV6), 1803, 1805, 1810 Human polyomavirus 7 (HPyV7), 1803, 1805, 1810 Human polyomavirus 9 (HPyV9), 1803, 1805, 1810 Human polyomavirus 12 (HPyV12), 1803, 1805, 1810 Human polyomaviruses, see Polyomaviruses Human rhinoviruses (HRVs), see Rhinoviruses Human T-cell lymphotropic viruses (HTLVs), 1458–1465 clinical significance, 1460–1461 collection and storage of specimens, 1406, 1461 description of agent, 1458 detection and identification methods, 1434 direct examination, 1462

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 92

epidemiology and transmission, 1458–1460 evaluation, interpretation, and reporting of results, 1464–1465 genomic organization, 1458–1459 PCR, 1461–1462 serologic testing algorithm, 1460 serologic tests, 1462–1464 taxonomy, 1458 treatment, 1461 typing systems, 1463–1464 vaccine, 1461 virus isolation and identification, 1462– 1463 Hutchinson’s teeth, 1059 HVISA (heterogeneous vancomycinintermediate S. aureus), 1230, 1295 H&V-Mix, 1426, 1427 FreshCells, 1413 virus susceptibility profiles, 1429 Hyaline, definition, 1941 Hyaline fungi, 2057–2077 antigen detection, 2076 antimicrobial susceptibilities, 2077 clinical significance, 2059, 2075–2076 collection, transport, and storage of specimens, 2059 description of agents, 2069–2075 direct examination, 2076 epidemiology and transmission, 2075 evaluation, interpretation, and reporting of results, 2077 identification, 2057–2059, 2069–2075, 2077 isolation, 2077 key phenotypic features, 2060–2065 microscopy, 2076 nucleic acid detection, 2076–2077 taxonomy, 2057–2059, 2069–2075 typing systems, 2077 Hyalohyphomycosis, 1947, 2057, 2076 Hyalomma, 2507, 2515 Hyaluronidase production, by Streptococcus, 395 Hybrid Capture 2 (hc2) test, 1786–1787, 1792 Hybrid capture assays, 56, 57 Hybridization allele-specific, 67, 69 dual hybridization probes for real-time PCR, 61 Hybridization arrays high-density arrays, 71–72 low- to moderate-density arrays, 72 Hybridization protection assay (HPA), HPV, 1793 Hybri-Max, 1791 Hydatid disease, 2330–2332 Hydrocele, filarial nematodes and, 2464– 2465 Hydrocortisone, 2515 Hydrogen peroxide, 194–195 catalase test, 317 Hydronyche, 2520 Hydrophobia, 1635 Hydrops fetalis, 1819–1820 Hydroxychloroquine, 2536–2537 adverse effects, 2537 mechanism of action, 2536 pharmacokinetics, 2537 spectrum of activity, 2537 Hydroxyquinoline, for Dientamoeba fragilis, 2413 Hydroxystilbamidine, for Acanthamoeba, 2394

Hygienic hand washing, 186–187 Hymenolepididae (family), 2291, 2475 Hymenolepis (genus), 2475, 2533 Hymenolepis diminuta, 2502 arthropod vector, 2507 detection, 2320 eggs, 2449, 2501 Hymenolepis nana, 2471–2472, 2475–2476, 2502 clinical significance, 2476 collection, transport, and storage of specimens, 2476 description, 2475 detection, 2320 direct examination, 2476 eggs, 2449 epidemiology, transmission, and prevention, 2475 evaluation, interpretation, and reporting of results, 2476 microscopy, 2476 serologic tests, 2476 taxonomy, 2475 treatment, 2476, 2531 Hymenoptera (order), 2518 Hyperestrogenism, 2190 Hyperkeratosis, phaeohyphomycoses and, 2162 Hypersensitivity, see also Allergy arthropods and, 2515 atypical measles syndrome, 1520 scabies, 2516 scorpion venom, 2520 to cetuximab, 2516 Hypersensitivity pneumonitis, Mycobacterium and, 599 Hyphae, 1935, 1937, 1939, 1941, 1966– 1967, 1969, 1973–1974 Hyphomyces destruens Hyphomycetes, 1939, 1941, 2058, 2063– 2065, 2071–2075 Hyplex MRSA, 361 Hyplex StaphyloResist, 361 Hyplex system, 1383–1384 Hypocalcemia, measles and, 1521 Hypochlorous acid, 195 Hypocreales (order), 1937, 1938, 2059, 2173–2174 Hypoderaeum, 2482 Hypoderma, 2330 Hypoderma lineatum, 2519 Hypoglycemia, Plasmodium falciparum, 2341 Hypostome, tick, 2512 Hypotension, spider envenomation and, 2520 Iam VZV Q-LAMP qualitative assay, 1707 IATA Dangerous Goods Regulations, 1416 Ibis system, 73 ICE Syphilis, 1068, 1071 ICEPlex system, 67 ICEPlex ViraQuant panel, 1726 ICTV, 1394 ID 32C, 2001, 2003 IDEIA Hp StAR, 1019 Identification of organisms, see also specific organisms aerobic Gram-positive cocci, 350–352 health care-associated infections, 110–111 methods, 259–263 amplified fragment length polymorphism (AFLP), 261 cell wall composition, 262 cellular fatty acid analysis, 262

SUBJECT INDEX chemical methods, 262 DNA-DNA hybridization studies, 259– 260 FTIR spectroscopy, 263 G+C content, 261 mass spectrometry, 262–263 PCR-based, 261 phenotypic methods, 261–262 rRNA studies, 260–261 sequence analysis of protein-encoding genes, 261 whole-genome sequence-based methods, 260 systems for bacteria and fungi, 29–40 evaluation of, 31 genotypic identification systems, 39–40 limitations of, 31–32 organism identification systems, 29–31 overview, 29–30 phenotypic identification systems, 32– 35 proteomic identification systems, 35–39 selection criteria, 31 system construction, 30–31 Idoxuridine, for herpes simplex virus (HSV), 1689 IgA testing, Toxoplasma gondii, 2377 IgE antibody detection, Toxoplasma gondii, 2377 IgG avidity assay, see Avidity assay IgM tests, 100 cytomegalovirus (CMV), 1729–1730 varicella-zoster virus, 1712 IgM capture ELISA arboviruses, 1653–1654 hantaviruses, 1664–1665 Toxoplasma gondii, 2376–2377 IgM-class antibodies, 94 Ignatzschineria larvae, 822 Ignavigranum clinical significance, 425 identification, 428, 429 taxonomy, 423 Ignavigranum ruoffiae, 423, 425 IHC, see Immunohistochemistry Ikoma lyssavirus, 1633–1634 Ileitis, Yersinia pseudotuberculosis, 742 Ileocolitis, measles, 1521 Ilesha virus, 1645 Ilheus virus, 1645 Illumigene mycoplasma assay, 1094 Illusory parasitosis, 2521 Imagen hMPV DFA test, 1503, 1511 Imagen influenza virus A and B, 1473 Imagen RSV, 1503 IMDx Flu A/B and RSV for Abbott m2000, 1477 Iminophenazine, 1360 Imipenem, 1176–1177 anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1255, 1260 Bacteroides fragilis group susceptibility percentages, 1350 concentration in serum, 1199 with MK7655, 1178 Imipenem resistance, 1347 Immersion fluid (immersion oil), 5, 8–9 Immulite 2000 Anti-HBc, 1848 Immulite 2000 HBc IgM, 1848 Immulite 2000 hepatitis B virus, 1847 Immulite 2000 Syphilis Screen, 1071 Immune reconstitution inflammatory syndrome (IRIS), 1161, 2018

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 93

Immunization, see Vaccine Immunoassays, 91–103; see also specific assay types agglutination, 96–97 automated, 102–103 categorization, 92 chemiluminescence immunoassay (CLA), 100–101 complement fixation, 97 definition of terms, 91–92 enzyme immunoassays (EIAs), 98–100 antibody interference, 100 competitive, 98 hook effect, 99–100 IgM measurement, 100 noncompetitive, 98, 99 plate variability, 99 technical challenges, 98–99 historical perspective, 91 human immunodeficiency virus, 1443– 1447 immunoblotting, 101 immunofluorescence assays, 97–98 immuno-PCR, 102–103 lateral-flow immunoassay (LFA), 102 multiplex, 102 neutralization assays, 97 parasites in stool specimens, 2319–2320, 2322 parvovirus B19, 1822 performance characteristics, 92–93 precipitation reactions, 95–96 protective immunity, analysis as measure of, 94 quantification, 95 rapid, 101–102 recent versus remote infection, analysis as measure of, 94–95 antibody titer, change in, 94 avidity testing, 94–95 IgM-class antibodies, 94 screening versus diagnostic assays, 93 sensitivity, 92 specificity, 92 specimen requirements, 94 Western blotting, 101 Immunoblot, 101 Anaplasma phagocyrophilum, 1143–1144 Borrelia, 1046–1047 Ehrlichia chaffeensis, 1142 Epstein-Barr virus, 1744, 1746 human herpesvirus 7 (HHV-7), 1762 Leishmania, 2361 parvovirus B19, 1822 schistosomes, 2486 Toxoplasma gondii, 2377 Treponema, 1070, 1072 ImmunoCAP assay, Aspergillus, 2044 ImmunoCard Mycoplasma pneumoniae, 1097– 1098 ImmunoCard STAT! assays for gastroenteritis viruses, 1624 ImmunoCard STAT! Campy assay, 1002 ImmunoCard STAT! Cryptosporidium/ Giardia, 2295 ImmunoCard STAT! Cryptosporidium/Giardia rapid assay, 2441 ImmunoCard STAT! Giardia duodenalis, 2412 ImmunoCard STAT! HpSA, 1019 ImmunoCard STAT! RSV Plus, 1504 Immunochromatographic assay Cryptosporidium, 2439, 2441 Epstein-Barr virus, 1743–1744

n xciii

gastroenteritis viruses, 1623–1624 Giardia duodenalis, 2411 malaria, 2335–2336 parasitology, 2295–2296 Trichomonas vaginalis, 2415 Immunocolorimetric assay (ICA), rubella virus, 1528–1530 Immunocompromised/immunosuppressed individual Acanthamoeba, 2391 adiaspiromycosis, 2115 antiviral susceptibility testing, 1914 Aspergillus, 2033, 2036 Balantidium coli, 2417 Candida, 1993 Coccidioides, 2114 Cryptococcus, 1993–1994 Cryptosporidium, 2437–2438 Cystoisospora belli, 2428 cytomegalovirus, 1719–1720 dermatophytoses, 2136 enteric adenoviruses, 1622 entomophthoromycosis, 2100 Epstein-Barr virus, 1739–1740 Fusarium, 2058, 2067 Giardia duodenalis, 2409 hepatitis E virus, 1590 herpes simplex virus (HSV), 1689 human herpesvirus 6 (HHV-6), 1754– 1756, 1760 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1763 human metapneumovirus, 1509 hyaline fungi, 2075–2076 Irpex lacteus, 2075 measles, 1521 microsporidia, 2210, 2213 Mycoplasma, 1092–1093 noroviruses, 1622 Paracoccidioides brasiliensis, 2115 parainfluenza virus, 1488 parvovirus B19, 1820, 1823 phaeohyphomycoses, 2162–2163 Pneumocystis, 2016–2019 polyomavirus, 1804–1806, 1812 respiratory syncytial virus (RSV), 1500– 1501 rotaviruses, 1622 scabies, 2516 Schizophyllum commune, 2075 Sporothrix, 2161 Strongyloides stercoralis, 2457–2458 Taenia crassiceps, 2477 Talaromyces marneffei, 2046 Toxoplasma gondii, 2374–2375, 2381–2382 Trypanosoma cruzi, 2362–2363 varicella-zoster virus, 1705–1706 Immunodiffusion (ID), 95–96 Blastomyces, 1969, 2120, 2123 Coccidioides, 1971, 2121, 2123 eumycotic mycetoma, 2181 fungi, 1969, 1971 Histoplasma capsulatum, 1971, 2120, 2122 Paracoccidioides brasiliensis, 2121, 2123 Pythium insidiosum, 2203 Immunoelectron microscopy, 1623, 1629 Immunofluorescence, 10 adenoviruses, 1773–1774, 1776 detection of Chlamydia and viruses, 1423– 1427 Herpesviridae, 1429 human metapneumovirus, 1511 mumps virus, 1494 parainfluenza virus, 1491–1492

xciv

n

SUBJECT INDEX

Immunofluorescence assay (IFA), 97–98 arboviruses, 1652–1654 Bartonella, 880 Borrelia, 1045–1046 Brucella, 867–868 Coxiella burnetii, 1154–1155 Cryptosporidium, 2439, 2441 cytomegalovirus (CMV), 1728–1729 Epstein-Barr virus, 1742, 1743–1745 hantaviruses, 1664–1665 human herpesvirus 6 (HHV-6), 1759 human herpesvirus 7 (HHV-7), 1759, 1762 human herpesvirus 8 (HHV-8), 1759, 1764 human T-cell lymphotropic viruses (HTLVs), 1462–1464 Legionella pneumophila, 898 measles virus, 1522–1523 Mycoplasma, 1097 Pneumocystis, 2023 respiratory syncytial virus, 1502, 1505 Rickettsia, 1124, 1128–1129 rubella virus, 1528–1530 Trichomonas vaginalis, 2415 Immunofluorescent antibody stain, 322–323 Immunoglobulin Epstein-Barr virus, 1740 for cytomegalovirus, 1720 for human metapneumovirus, 1509 hepatitis B (HBIG), 1844 human metapneumovirus, 1512 polyomavirus-associated nephropathy (PVAN), 1811 vaccinia, 1831–1832 Immunohistochemistry (IHC) arenaviruses, 1676 Brachyspira, 1064 Coxiella burnetii, 1153 Epstein-Barr virus antigen detection by, 1742 filoviruses, 1676 Francisella tularensis, 856 hantaviruses, 1663 human herpesvirus 6 (HHV-6), 1756, 1759 human herpesvirus 7 (HHV-7), 1759 human herpesvirus 8 (HHV-8), 1759, 1763 human papillomaviruses (HPV), 1788– 1789 parvovirus B19, 1821 polyomavirus, 1806 poxviruses, 1832 Treponema pallidum, 1062–1063, 1073 Tropheryma whipplei, 1162 Immunologic methods for culture confirmation of Neisseria, 641 ImmunoWELL Mycoplasma pneumoniae IgM and IgG, 1098 IMMY cryptococcal lateral-flow assay, 1997 IMP type β-lactamases, 1223–1224, 1226, 1383–1384 Imperial College London, 150 Impermeability mutations, in Pseudomonas aeruginosa, 781 Impetigo, Streptococcus pyogenes and, 385 In situ hybridization, 54, 55 human herpesvirus 6 (HHV-6), 1756 PNA fluorescent, 55 Incomplete metamorphosis, 2508, 2510– 2511, 2513, 2520 Index of refraction, 6 India ink, 1957, 1970, 1975, 1988, 1996

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 94

Indicators, 320, 321 Indinavir, for human immunodeficiency virus (HIV), 1871, 1875–1876 Indinavir resistance, 1897–1898 Indirect fluorescent antibody (IFA) test, 97 Acanthamoeba, 2394 Anaplasma phagocyrophilum, 1143–1144 arenaviruses, 1676, 1679–1680 Babesia, 2352 coronaviruses, 1570, 1577 cytomegalovirus (CMV), 1728–1729 detection of Chlamydiae and viruses, 1423–1425 Ehrlichia chaffeensis, 1140, 1142 filoviruses, 1676, 1679–1680 human immunodeficiency virus, 1446– 1447 influenza viruses, 1472–1473 Leishmania, 2361 mumps virus, 1495 parainfluenza virus, 1489, 1491 Plasmodium, 2349 rabies virus, 1640–1641 Toxoplasma gondii, 2376–2380 Trypanosoma brucei, 2367 Trypanosoma cruzi, 2365 Indirect hemagglutination Entamoeba histolytica, 2405 Trypanosoma brucei, 2367 Trypanosoma cruzi, 2365 Indirect rapid immunohistochemistry test (IRIT), rabies virus, 1640 Indole test, 318, 615–616 Inermicapsifer madagascariensis, 2502 Infant botulism, 947 Infection prevention committee, 108–109 Infection prevention program (IPP), 106– 110, 112 Infectious bronchitis virus (IBV), 1566 Infectious diarrhea, etiologies of, 290 Infectious disease syndromes, usual etiologies of, 290–291 Infectious mononucleosis, 1738–1740 Infertility Chlamydia trachomatis, 1108 Mycoplasma, 1092 Ureaplasma, 1092 Infiniti analyzer, 72 Infiniti HPV, 1791 Infiniti HPV-HR Quad, 1791 Infiniti HPV-Quad, 1791 Infiniti respiratory viral panel, 1506, 1511 Infiniti RVP Plus, 1478, 1555, 1576 Inflammatory bowel disease (IBD) colonic microbiome in, 230–231 Helicobacter, 1018 Yersinia enterocolitica, 742 Influenza antiviral susceptibility testing, 1916, 1921, 1924 clinical significance, 1470–1471 epidemiology and transmission, 1470–1471 HPAI (highly pathogenic avian influenza), 1481–1482 influenza A infection, 1470–1471 influenza B infection, 1470–1471 influenza C infection, 1471 laboratory tests suggested for, 125 laboratory-acquired infections, 177 pandemics, 1470–1471 treatment, 1471, 1903 vaccines, 1471–1472 Influenza A Duet stain, 1427 Influenza viruses, 1470–1482

antigen detection, 1472–1476 antiviral agents, 1886–1887 M2 protein inhibitors, 1886–1887 neuraminidase inhibitors, 1887 table of agents, 1886 antiviral resistance, 1903–1905, 1917 M2 channel blocker resistance, 1903– 1904 neuraminidase inhibitor resistance, 1903–1905 antiviral susceptibilities, 1481–1482, 1903 antiviral susceptibility testing, 1916 avian strains, 1470–1471, 1481–1482 biosafety, 1479, 1482 cell culture, 1476, 1479–1480 clinical significance, 1471–1472 cytopathic effect (CPE), 1479 description of agents, 1470 detection and identification methods, 1434 DFA and IFA reagents for the detection of, 1425 direct detection, 1472–1476 electron microscopy, 1472 epidemiology and transmission, 1470–1471 evaluation, interpretation, and reporting of results, 1482 genetic reassortment, 1470 genome sequencing, 246 H1N1, 1432, 1470–1472, 1481–1482, 1903 H2N2, 1470 H3N2, 1470–1472, 1481–1482, 1903 H5N1, 1470–1472, 1476, 1481–1482, 1903 H7N7, 1471 H7N9, 1470–1471, 1903 HPAI (highly pathogenic avian influenza), 1481–1482 identification and typing systems, 1480– 1481 immunofluorescence detection in R-Mix cells, 1426 isolation from embryonated chicken eggs, 1480 isolation procedures, 1476, 1479–1480 microscopy, 1472 naming of virus strains, 1470 nucleic acid analyses, 1476–1478 rapid cell culture, 1426 serologic tests, 1481 specimen collection and handling, 1406– 1408, 1472 taxonomy, 1470 transport medium for, 1409 vaccines, 1471–1472 Influenzavirus A (genus), 1398, 1470 Influenzavirus B (genus), 1398, 1470 Influenzavirus C (genus), 1398, 1470 Inform HPV II Family 6 probe, 1789 inhA gene, 1357–1358, 1358, 1360 Inhibitory mold agar, 1952, 1961, 2139 Innate immune system, herpes simplex virus (HSV) and, 1687 INNO-LIA HIV I/II Score test, 1446 INNO-LIA Syphilis score, 1070, 1072 INNO-LiPA, 1791, 1795 INNO-LiPA HBV DR version 2, 1921 INNO-LiPA HBV genotyping kit, 1921 INNO-LiPA-Mycobacteria v2 assay, 575, 582–583, 601 Innova, 48 Inonotus, 2062 Inonotus tropicalis, 2062, 2071, 2073, 2075

SUBJECT INDEX InoqulA FA/MI, 48 InPouch TV, 2315, 2327 Inquilinus limosus, 626–627, 822–823 Insecta (class), 2507, 2522–2523 Instars, 2505 InSTEDD (Innovative Support to Emergencies, Disease, and Disasters), 128 INSTI HIV-1 antibody test kit, 1445 Institut Pasteur, 150 Integrase inhibitor(s), 1440 Integrase inhibitor resistance, 1447, 1897– 1899 Integrase strand transfer inhibitors, 1872, 1877–1878 Integrons Aeromonas, 1326 tet gene containing, 1234 Interferon alpha hepatitis B virus, 1881–1882, 1900 hepatitis C virus, 1601–1602, 1609–1611, 1878–1879, 1901 hepatitis E virus, 1591 human herpesvirus 8 (HHV-8), 1763 human T-cell lymphotropic viruses (HTLVs), 1461 polyomavirus, 1811 progressive multifocal leukoencephalopathy (PML), 1811 rabies virus, 1641 Interferon beta, for HTLVs, 1461 Interferon gamma release assays (IGRAs), 546, 555–556, 576 Interferon resistance hepatitis B virus, 1851, 1900 hepatitis C virus, 1901–1902 International Air Transport Association (IATA), 178, 1416, 1636, 1663 International Civil Aviation Organization (ICAO), 178 International Code of Nomenclature for algae, fungi and plants (ICN), 1936 International Code of Nomenclature of Bacteria, 263 International Code of Nomenclature of Prokaryotes, 258 International Code of Virus Classification and Nomenclature, 1394 International Code of Zoological Nomenclature, 2286 International Committee on Taxonomy of Viruses (ICTV), 1393–1394 character list, 1394–1395 database, 1402 website, 1394 International Journal of Systematic and Evolutionary Microbiology, 264 International Nucleotide Sequence Database Collaboration (INSDC) database, 580 International Organization for Standardization (ISO), 170, 183, 1249, 1253, 1268 International Society of Travel Medicine (ISTM), 128 International Working Group on the Classification of Staphylococcal Cassette Chromosome Elements, 139 InterProScan, 233 Interstitial pneumonitis, arenavirus, 1674 Intertrigo, Fusarium and, 2058 Intestinal obstruction, see Bowel obstruction Intestinal schistosomiasis, 2480 Intestinal tract specimen, see also Stool specimen

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 95

duodenal contents, 2325–2326 parasitology, 2294, 2298–2299, 2304– 2305, 2324–2326 pinworm examination, 2324 recovery of parasites, 2325 sigmoidoscopy material, 2324–2325 Intestinal trematodes, 2482, 2484, 2490 Intra-abdominal infection, Citrobacter and, 720 Intrasporangiaceae, 478 Intrauterine contraceptive device (IUD) Actinomyces, 923 non-spore-forming, anaerobic, Grampositive rods, 923 specimen collection, transport, and handling, 297 Intravenous drug users, HTLVs and, 1459, 1465 Intussusception, adenoviruses and, 1772 Invader assays, 56–57, 58, 1606 Invasive meningococcal disease, 636, 637, 643, 644 Investigational-use-only (IUO) products, 82 Iodamoeba, 2399–2400 Iodamoeba bütschlii, 2400, 2408 detection, 2317–2318, 2321 trophozoites and cysts, 2401–2402 Iodine stain, 2313 Iodophors, 185–186 Iodoquinol, 2541–2542 Balantidium coli, 2417 Blastocystis hominis, 2407 Dientamoeba fragilis, 2413 Entamoeba histolytica, 2405 Ion sequencing, 70–71 Ion Torrent system, 70–71 IPlex MassArray system, 73 Ippy virus, 1669, 1671 Iridocyclitis, Onchocerca volvulus and, 2466 IRIS (immune reconstitution inflammatory syndrome), 1161, 2018 Irkut virus, 1633–1634 Iron deficiency anemia, 2456 Iron-hematoxylin stain, 2314, 2318–2319 Iron-hematoxylin stain/carbol fuchsin, 2318– 2319 Irpex lacteus, 2071, 2075 IS1245/IS900, 585 IS6110 fingerprinting, 136–137 IS6110 RFLP typing, 584 Isavuconazole, 2230 dimorphic fungi, 2122 eumycotic mycetoma fungi, 2182 IsoAmp HSV assay, 1691 Isolate, definition, 132 Isolation procedures, see specific organisms Isolator blood culture system, 20 Isolator system, 293 Isoniazid activity, 1357 adverse effects, 1357 antimicrobial susceptibility testing, 1365– 1367 for Mycobacterium infection, 1357–1358 Isoniazid resistance, 1356–1358, 1363, 1367 Isonicotinic acid, 1360 Isoptera (order), 2522 Isosatratoxins, 2190 Iso-Sensitest agar, 1268 Isospora belli, 1192, 2425; see also Cystoisospora belli Isothermal amplification assays, for noroviruses, 1626–1627 IsoVitaleX, 672

n xcv

Isoxazolyl penicillins, 1171 Issatchenkia, 1937, 1938 Isthmiophora, 2482 Itch, arthropods and, 2515 Itching, see Pruritus Itraconazole, 2225–2226 antifungal susceptibility testing, 2255– 2273 Aspergillus, 2044–2045 chromoblastomycosis, 2167 dermatophytes, 2145 dimorphic fungi, 2121–2122 eumycotic mycetoma fungi, 2181–2182 Fusarium, 2069 hyaline fungi, 2077 mucormycosis, 2089 phaeohyphomycosis, 2167 spectrum of activity, 2224, 2225 Talaromyces marneffei, 2048 Trypanosoma cruzi, 2365 yeast species, MICs for, 2005 Itraconazole resistance, 2225, 2238–2239, 2241 ITS 1 sequence, 580 IVD Cryptosporidium antigen detection assay, 2441 IVD Research Giardia/Crypto Combo EIA, 2441 Ivermectin, 2533–2534 adverse effects, 2534 filarial nematodes, 2465 Gnathostoma, 2498 indications for, 2534 lice, 2511 Loa loa, 2468 Mansonella ozzardi, 2468 mechanism of action, 2533 Onchocerca volvulus, 2467 pharmacokinetics, 2533–2534 spectrum of activity, 2534 Strongyloides stercoralis, 2455, 2458 Ivory Coast ebolavirus, 1670 Ixodes, 1037–1040, 1138, 2507, 2514, 2516, 2523 Ixodes cookei, 2515 Ixodes dammini, 2512, 2515, 2523 Ixodes dentatus, 2515 Ixodes granulatus, 2515 Ixodes holocyclus, 2515–2516 Ixodes ovatus, 2515 Ixodes pacificus, 2515 Ixodes persulcatus, 2515 Ixodes ricinus, 2512, 2515–2516, 2523 Ixodes scapularis, 2512, 2515 Ixodida (order), 2511 Ixodidae (family), 2512, 2514 Janibacter chemotaxonomic features, 475 description of genus, 478 identification, 495–496 Janssen, Hans, 7 Janssen, Zaccharias, 7 Japanese encephalitis virus, 125, 1644–1645, 1647–1648, 1650, 1652–1655 Japanese Society for Chemotherapy, 1268 Japanese spotted fever, 1125 Jarisch-Herxheimer reaction, 1041 Jaundice, arbovirus, 1647 JC polyomavirus (JC virus), 1803–1812 antigen detection, 1806–1807 antiviral susceptibilities, 1811 cell culture, 1810 clinical significance, 1804, 1806

xcvi

n

SUBJECT INDEX

JC polyomavirus (JC virus) (continued) cytopathic effect (CPE), 1810 description of agents, 1803–1804 direct examination, 1806–1810 epidemiology and transmission, 1804 evaluation, interpretation, and reporting of results, 1812 in situ hybridization (ISH), 1807–1808 isolation procedures, 1810 microscopy, 1808 nucleic acid amplification tests (NAATs), 1807, 1809–1810 nucleic acid detection, 1807, 1809–1810 serologic tests, 1810 specimen collection and handling, 1407, 1806 taxonomy, 1803 JC polyomavirus (species), 1803 JC Virus r-gene Primers/Probe, 1810 JC/BK Consensus, 1810 Jeotgalicoccus, 354, 356–357, 361 Jigger, 2509–2510 Joint Commission on Accreditation of Healthcare Organizations, 203 Joint fluid specimen collection, transport, and storage guidelines, 276 fungi, 1946–1947, 1950 Joint infection Aggregatibacter, 654 anaerobic Gram-negative rods, 972 Brucella, 865 Finegoldia magna, 910–911, 911 Fusobacterium, 973 Mycobacterium, 599 non-spore-forming, anaerobic, Grampositive rods, 923 parvovirus B19, 1819 prosthetic joint Helcococcus, 425 Lactococcus, 424 Propionibacterium acnes, 924 Tropheryma whipplei, 1159–1160 Joint pain, spider envenomation and, 2520 Jonquetella anthropi, 968, 974, 980–981 Josamycin, 1182 Junin virus, 1669, 1671, 1673–1681 Kaffir virus, 1830 Kala-azar, 2358 Kanamycin, 1181–1182, 1220, 1358–1360 Kanamycin resistance, 1356, 1360 Kanamycin-esculin-azide agar, 338 Kandleria vitulina, 920 Kaposi’s sarcoma, 240, 1762–1764 Kaposi’s sarcoma-associated herpesvirus, see Human herpesvirus 8 Katayama fever, 2484 katG gene, 1357–1358 Kato-Katz method, 2474–2475, 2486 Kawasaki disease, 1773 Kazachstania pintolopesii, 2000 KCRYP, 2441 Kerandel’s sign, 2366 Keratitis Acanthamoeba, 2330, 2387, 2391, 2394 Acremonium, 1949 Aeromonas, 754 Aspergillus, 1949 Bacillus cereus, 443 Bacillus megaterium, 443 Beauveria bassiana, 2076 Candida albicans, 1949 Candida parapsilosis, 1949

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 96

Candida tropicalis, 1949 Capnocytophaga, 654 Curvularia, 1949 etiologies, usual, 290 fungal, 1949 Fusarium, 1949, 2058, 2067 herpes simplex virus (HSV), 1689 hyaline fungi, 2076 Lagenidium, 2198, 2204 Metarhizium anisopliae, 2076 Moraxella, 814 Mycobacterium, 599–600 Nocardia, 515 Nocardia farcinica, 517 Onchocerca volvulus, 2466 Pseudomonas aeruginosa, 775 Pythium insidiosum, 2198, 2201 Scopulariopsis brevicaulis, 2075 Tsukamurella tyrosinosolvens, 519 viruses, specimens and methods for detection of, 1407 Keratoconjunctivitis adenoviruses, 1772 microsporidia, 2210, 2213, 2216 Kernia, 2071 Kerstersia antimicrobial susceptibilities, 845 clinical significance, 841 collection, transport, and storage of specimens, 842 description of genus, 839 evaluation, interpretation, and reporting of results, 845 identification, 843 taxonomy, 838 Kerstersia gyiorum, 614, 838, 841, 843 Kerstersia similis, 838, 843 Ketamine, for rabies virus, 1641 Ketoconazole, 2226 antifungal susceptibility testing, 2255– 2273 eumycotic mycetoma fungi, 2181–2182 Leishmania, 2361 Pythium insidiosum, 2203 spectrum of activity, 2226 Talaromyces marneffei, 2048 Trypanosoma cruzi, 2365 Ketoconazole resistance, 2226 Ketolide(s), 1184–1185 adverse effects, 1185 mechanism of action, 1184 pharmacology, 1184 spectrum of activity, 1184 Ketolide resistance, 1184, 1231 KHCRYP, 2441 Khujand virus, 1633–1634, 1640 KI polyomavirus (KIPyV), 1803–1805, 1810 Kickxellales (order), 2087 Kickxellomycotina (subphylum), 2087 Kidney disease/failure Corynebacterium diphtheriae, 480 Plasmodium falciparum, 2341 Kidney specimen, for parasitology, 2328, 2330 Kinetoplastida (class), 2287 Kingella antimicrobial susceptibilities, 662 antimicrobial susceptibility testing, 1328 clinical significance, 655 direct examination, 656 epidemiology and transmission, 653 identification, 660–661 isolation procedures, 656 taxonomy and description of, 653

Kingella denitrificans, 653, 655, 660–661 Kingella indologenes, see Suttonella indologenes Kingella kingae, 653, 655–657, 660–661 Kingella oralis, 653, 655, 661 Kingella potus, 653, 655, 661 Kinyoun acid-fast stain, 321–322, 1957, 2313 Kirby-Bauer method, 1269 Kissing bugs, 2508, 2522 Klebsiella antimicrobial susceptibilities, 727–730, 1173–1174, 1178, 1186, 1195– 1196 antimicrobial susceptibility testing, 1279 β-lactamases, 1299 description of genus, 715 epidemiology, transmission, and clinical significance, 715–716, 718 evaluation, interpretation, and reporting of results, 731 identification, 723–725 Klebsiella granulomatis, 715, 722, 725 Klebsiella oxytoca, 716, 718, 724, 727 antibiotic resistance, 1226–1227 antimicrobial susceptibility testing, 1279 Klebsiella ozaenae, 716, 718, 723–724 Klebsiella pneumoniae, 714–716 antibiotic resistance, 1212, 1217––1218, 1220, 1223–1226, 1228, 1232, 1234 antimicrobial susceptibilities, 727–728, 1177, 1187 antimicrobial susceptibility testing, 1279 as ESKAPE pathogen, 714 β-lactamases, 1223–1226, 1228, 1299 endoscope contamination outbreak, 199 epidemiology, transmission, and clinical significance, 715–716, 718 evaluation, interpretation, and reporting of results, 731 identification, 723–725 reference strains, 1299 typing systems, 727 Klebsiella pneumoniae carbapenemase, 695– 696 Klebsiella pneumoniae subsp. granulomatis, 716, 718 Klebsiella pneumoniae subsp. pneumoniae, 716 Klebsiella rhinoscleromatis, 716, 718, 723–724 Klebsiella singaporensis, 716 Klebsiella variicola, 716, 718, 725 Kligler iron agar, 338 Kluyvera, 722, 725–726, 1225 Kluyvera ascorbata, 718, 722 Kluyvera cryocrescens, 718, 722 Kluyvera georgiana, 718 Kluyvera intermedia, 718 Kluyveromyces, 1937, 1938 Kluyveromyces marxianus, 1985, 2000 Knoellia chemotaxonomic features, 475 description of genus, 478 identification, 495–496 Knott concentration, 2307, 2336, 2465 Knufia, 2153, 2154, 2158 Knufia epidermidis, 2154, 2158, 2162 Kobuvirus (genus), 1618 Koch, Robert, 238, 246, 444 Koch’s postulates, 238, 246 Koch-Weeks bacillus, 670 Kocuria, 354, 356, 361 Kocuria carniphila, 367 Kocuria kristinae, 355–356, 366–367 Kocuria rhizophila, 361, 367

SUBJECT INDEX Kocuria rosea, 356, 366–367 Kocuria varians, 356–357, 366–367 Kodamaea, 1988 Kodamaea ohmeri, 1985, 1993, 2005 Kodo poisoning, 2189 Kodoko virus, 1671 Köhler, August, 5 Köhler illumination, 5 Koplik’s spots, 1520 Kosakonia, 715, 719, 723 Kosakonia arachidis, 716, 719 Kosakonia cowanii, 716, 719, 723 Kosakonia oryzae, 716, 719 Kosakonia radicincitans, 716, 719, 723 Kovac’s reagent, 318 KPC β-lactamases, 1225, 1279, 1300, 1383– 1384 Kroppenstedtia chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 515 description of genus, 506 morphologic characteristics, 507 taxonomy, 504–505 Kroppenstedtia eburnea, 506, 514–515 Kujo virus, 243 Kunjin virus, 1645 Kuru, 1859 Kwell, 2517 Kyasanur Forest virus, 1645 Kytococcus, 354, 356, 361 Kytococcus aerolatus, 356 Kytococcus schroeteri, 356, 367 Kytococcus sedentarius, 356, 367 L forms, 1088 Label-free assays, 92 Labeling, specimen, 283–284 LabNet, WHO, 1523–1524 Laboratory biosafety, 169–178 audits, 170 clinical laboratory design, 171–172 MALDI-TOF MS and, 178 notable laboratory-acquired infections, 176–178 personal attributes, 170–171 point-of-care testing (POCT), 178 post-exposure management, 178 program, 169–170 quality management, 174–176 gloves, 175–176 hand washing, 175 immunization, 176 material safety data sheets (MSDS), 175 personal protective equipment, 175 respiratory masks, 176 safety preparedness, 174–175 risk management, 170 risk-based classification of microorganisms, 171 safety equipment, 172–174 biosafety cabinets, 172, 173 centrifuges, 174 chemical fume protection, 172–173 installation, 173 medical waste, 174 sharps protection, 174 splashguards, 172 standard practices, 173 transportation of samples, 178 Laboratory detection of bacteremia and fungemia, 15–26 assessment of methods for detection, 15

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 97

critical factors in pathogen recovery, 16– 17 agitation of culture bottles, 17 anticoagulant, 17 dilution of blood, 17 medium and additives, 17 number of cultures, 16–17 volume of blood cultured, 16 culture-based methods, 19–22 automated systems, 20–22 BacT/Alert, 21, 22 Bactec 9000 series, 21, 22 lysis-centrifugation system, 20 manual blood culture systems, 20 pediatric blood culture bottles, 22 VersaTREK, 21–22, 23 diagnostic importance, 15 interpretation of blood culture results, 18– 19 non-culture-based methods, 22–23 nucleic acid amplification, 23 surrogate markers for sepsis, 22–23 prognostic importance, 16 quality audits and benchmarks, 25 rapid identification of microbial isolates, 23–25 antigen detection, 24 direct rapid antimicrobial susceptibility testing from blood culture bottles, 25 MALDI-TOF (MS), 24–25 multiplex technology, 24–25 nucleic acid amplification test, 24 peptide nucleic acid-fluorescent in situ hybridization (PNA-FISH), 23 specimen collection, 17–18 number and timing of cultures, 18 skin disinfection, 17 Laboratory handling of specimens, 283–286 autopsy samples, 305–306 blood cultures, 292–294 cerebrospinal fluid, 294 culture examination and interpretation, 289 documentation, 283–284 ears, 295 eyes, 295 gastrointestinal tract, 301–303 beta-hemolytic streptococci, 303 Clostridium botulinum, 303 food poisoning, 302 Helicobacter pylori, 303 MRSA, 302–303 Shiga toxin-producing E. coli (STEC), 302 small bowel bacterial overgrowth syndrome, 303 VRE, 303 genital tract, 295–297 Actinomyces, 297 bacterial vaginosis (BV), 296 C. trachomatis, 295–296 dark-field examination for T. pallidum, 296–297 group B streptococcus screening, 296 H. ducreyi, 297 N. gonorrhoeae, 295–296 vaginitis, 296 Gram stain, 285–288 initial sample handling, 285–289 labeling of specimens, 283–284 lower respiratory tract, 297–299 bronchoscopy, 298 Chlamydia and Chlamydophila, 299

n xcvii

cystic fibrosis patients, 299 Legionella, 298 M. pneumoniae, 298–299 Nocardia, 299 medium inoculation, 286–287, 289 molecular detection of bacteria, 289 prioritization, 285 processed at remote site, 285 rejection of specimens, 284–285 reporting results, 289 tissue biopsy samples, 289–292 bone marrow, 291 cellulitis, 292 lymph nodes, 291–292 necrotizing fasciitis, 292 placenta, 292 quantitative culture, 292 uncultivable bacteria, 292 upper respiratory tract, 299–301 A. haemolyticum, 300 C. diphtheriae, 300–301 epiglottitis, 301 nasal, 300 nasopharynx, 300 sinus, 300 throat, 299–300 urinary tract, 303–305 bacterial antigen testing, 305 culture, 303–304 leptospires, 304–305 wounds, 305 Laboratory information system (LIS) antimicrobial susceptibility testing, 1274– 1277, 1281 Laboratory Preparedness Exercise (LPX) proficiency test, 219 Laboratory Response Network (LRN), 217– 219, 446, 448 Laboratory safety issues regarding bacterial pathogens, 282– 283 education and a culture of safety, 283 engineering controls, 282–283 personal protective equipment, 283 Mycobacterium and, 544, 546–548 Laboratory-acquired infections epidemiology, 169 external quality assessment and, 177–178 microorganisms associated with, 176–178 Brucella, 176 Burkholderia pseudomallei, 176–177 Coccidioides, 177 Francisella tularensis, 176 influenza, 177 Mycobacterium tuberculosis, 177 prions, 177–178 viral infections, 177 prevention, see Laboratory biosafety Laboratory-developed tests (LDTs), 79–80, 82, 1655 LabPro, 32, 1275, 1277 LabPro Alert, 1277 LabPro with AlertEX, 33 Lacazia loboi, 2110, 2196–2199 antimicrobial susceptibilities, 2199 clinical significance, 2197–2198 collection, transport, and storage of specimens, 2198 description, 2196 direct examination, 2198 epidemiology and transmission, 2196–2197 evaluation, interpretation, and reporting of results, 2199 identification, 2199

xcviii n

SUBJECT INDEX

Lacazia loboi (continued) microscopy, 2198–2199 phylogeny, 2197–2198 serologic tests, 2199 taxonomy, 2196, 2198 Lacaziosis, 2196–2199 Lachnoanaerobaculum saburreum, 922, 924, 930 Lachnospira, 922 Lachnospira pectinoschiza, 922 Lachnospiraceae (family), 922, 924 LA-Cocci antibody system, 2121 La Crosse virus, 1644–1645, 1647, 1651 Lactic acidosis, 1191 Lactobacillaceae (family), 920 Lactobacillales (order), 354, 441, 920 Lactobacilli, in vaginal microbiome, 231 Lactobacillus antibiotic resistance, 1234 antimicrobial susceptibilities, 931, 1175, 1183–1184, 1189–1190, 1348 antimicrobial susceptibility testing, 1254, 1317, 1329–1330 clinical significance, 924 identification, 438, 927, 930 in vaginal microbiome, 231 isolation procedures, 926 lack of in bacterial vaginosis, 906 taxonomy and description, 920–921 Lactobacillus acidophilus, 924, 927, 1329, 1348 Lactobacillus casei, 924, 931, 1348 Lactobacillus catenaformis, see Eggerthia catenaformis Lactobacillus confusus, 424 Lactobacillus crispatus, 924, 1348 Lactobacillus delbrueckii, 1329 Lactobacillus fermentum, 924, 1348 Lactobacillus gasseri, 924, 1329, 1348 Lactobacillus iners, 618, 924, 926 Lactobacillus jensenii, 924, 1348 Lactobacillus johnsonii, 1348 Lactobacillus plantarum, 924, 1348 Lactobacillus rhamnosus, 924, 927, 931, 1329, 1348 Lactobacillus salivarius, 1348 Lactobacillus uli, see Olsenella uli Lactobacillus ultunensis, 924 Lactobacillus vitulinus, see Kandleria vitulina Lactococcus antimicrobial susceptibilities, 430 clinical significance, 424 epidemiology and transmission, 423 identification, 425–426, 426, 429 taxonomy, 422–423 Lactococcus garvieae, 422, 424, 429, 430 Lactococcus lactis, 422, 429, 430 Lactococcus mesenteroides, 429 Lactophenol cotton blue, 1958 Lactrimel agar (Borelli’s medium), 1961 Lady Windermere syndrome, 541 Lagenidiaceae (family), 2203 Lagenidiales (order), 2203 Lagenidiosis, 2203–2204 Lagenidium, 1936, 1939, 2203–2204 antimicrobial susceptibilities, 2204 clinical significance, 2198, 2204 description, 2203–2204 direct examination, 2204 epidemiology and transmission, 2198, 2204 evaluation, interpretation, and reporting of results, 2204 microscopy, 2202, 2204 phylogeny, 2197–2198

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 98

serologic tests, 2204 taxonomy, 2198, 2203 Lagos bat virus, 1633–1634 Lagovirus (genus), 1617 Laguna Negra virus, 1661 L-Alanine-7-amido-4-methycoumarin, 316 Lamivudine hepatitis B virus, 1881–1882, 1900 human immunodeficiency virus (HIV), 1870, 1872–1873 human T-cell lymphotropic viruses (HTLVs), 1461 Lamivudine resistance, 1917, 1921 hepatitis B virus, 1851, 1900 HIV, 1896–1898 LAMP, see Loop-mediated amplification Lancefield M protein, in serotyping, 139 Lantern cells, 1821 LAP test (leucine naphthylamide), 318–319 Larva currens, 2457 Laryngeal HPV, 1786 Laryngospasm, spider envenomation and, 2520 Laryngotracheobronchitis, measles, 1521 Lash’s casein hydrolysate-serum medium, 2315 Lasiodiplodia, 2153 Lasiodiplodia theobromae, 2153–2154, 2156, 2162–2163 Lassa virus, 1647 antiviral susceptibilities, 1681 clinical significance, 1673–1674 collection, transport, and storage of specimens, 1674–1675 epidemiology and transmission, 1670 identification of virus, 1679–1680 isolation procedures, 1678 laboratory tests suggested for, 125 nucleic acid detection, 1677 serologic diagnosis, 1680–1681 taxonomy, 1669, 1671 Latency Epstein-Barr virus, 1738 herpes simplex virus (HSV), 1688 human herpesvirus 6 (HHV-6), 1754 human herpesvirus 8 (HHV-8), 1762, 1764 human immunodeficiency virus clinical latency, 1439 Lateral-flow assays, 102 Aspergillus, 2038 Cryptosporidium, 2439, 2441 Lateral-flow device (LFD), 1972, 1977 Lateral-flow immunochromatography, for adenoviruses, 1773–1774, 1776 Lateral gene transfer (LGT), 255, 257–258 Latex agglutination test adenovirus, 1774, 1776 Coccidioides, 2121, 2123 Cryptococcus, 1977 Entamoeba histolytica, 2405 Epstein-Barr virus, 1743–1744 gastroenteritis viruses, 1623–1624 Histoplasma capsulatum, 2120, 2122 rubella virus, 1529 Sporothrix schenckii, 2167 Trichomonas vaginalis, 2415 Latex-Cryptococcus antigen test, 1997 LATEX/T. b. gambiense, 2367 Latino virus, 1669, 1671 Latrine fly, 2519 Latrodectism, 2520 Latrodectus, 2520 Lawrence Livermore microbial detection array, 241

LCMV, 1407 Lean management method, 47 Lecithinase, for Clostridium identification, 953–954 Lecithodendriidae (family), 2290 Leclercia, 722, 726 Leclercia adecarboxylata, 715, 718, 722, 725 Lecythophora, 2069, 2071 Lecythophora hoffmannii, 2071 Lecythophora mutabilis, 2071 Ledipasvir, for hepatitis C virus infection, 1601, 1879–1880 Ledipasvir resistance, 1902–1903 LEE (locus for enterocyte effacement) pathogenicity island, Escherichia coli, 686, 688–689, 694–695 Leeming and Notman medium, 1961 Leeming’s medium, 1953 Leflunomide herpesviruses, 1886 polyomavirus, 1811 polyomavirus-associated nephropathy (PVAN), 1811 progressive multifocal leukoencephalopathy (PML), 1811 Leg weakness, in tick paralysis, 2516 Legionella, 887–899 antibody determination, 898 antigen detection, 891–892 antimicrobial susceptibilities, 898–899, 1177, 1180, 1183–1184, 1187, 1195 antimicrobial susceptibility testing, 898– 899 characteristics of species, 888 clinical significance, 890 collection, storage, and transport, 298, 890 description of genus, 887, 889 direct examination, 890–892 epidemiology, transmission, and pathogenesis, 889–890 evaluation, interpretation, and reporting of results, 899 identification from bacteria colonies, 895– 898 advanced identification, 897–898 basic identification, 895, 897 flow scheme, 896 in Acanthamoeba, 2389 isolation procedures, 892–895 culture media, 893–894 initial workup of suspect colonies and look-alike bacteria, 895 inoculation of plates, 893 medium incubation, 894 plate inspection, 894 specimen plating, 892–893 microscopy, 890–891 molecular diagnosis Legionnaires’ disease, 892 taxonomy, 887 typing systems, 898 Legionella adelaidensis, 888 Legionella anisa, 888, 2389 Legionella beliardensis, 888 Legionella birminghamensis, 888 Legionella bozemanae, 888, 899 Legionella brunensis, 888 Legionella cardica, 888 Legionella cherrii, 888 Legionella cincinnatiensis, 888 Legionella drancourtii, 888 Legionella dresdenensis, 888 Legionella drozanskii, 888

SUBJECT INDEX Legionella dumoffii, 887–888, 899 Legionella erythra, 888 Legionella fairfieldensis, 888 Legionella fallonii, 888 Legionella feeleii, 888 Legionella geestiana, 888 Legionella gormanii, 888 Legionella gratiana, 888 Legionella gresilensis, 888 Legionella hackeliae, 888 Legionella impletisoli, 888 Legionella israelensis, 888 Legionella jamestownensis, 888 Legionella jordanis, 888, 895 Legionella lansingensis, 888 Legionella londiniensis, 888 Legionella longbeachae, 887–889, 893, 899 Legionella lytica, 888 Legionella maceachernii, 888 Legionella massiliensis, 888 Legionella micdadei, 321, 550, 887–888, 891, 893, 899 Legionella moravica, 888 Legionella nagasakiensis, 888, 895 Legionella nautarum, 888 Legionella oakridgensis, 888, 894–895 Legionella parisiensis, 888 Legionella pneumophila, 887–899 antibody determination, 898 antigen detection, 891 antimicrobial susceptibilities, 898–899, 1190, 1195 antimicrobial susceptibility testing, 898– 899 characteristics of species, 888 clinical significance, 890 description of agent, 887, 889 direct examination, 890–892 epidemiology, transmission, and pathogenesis, 889–890 evaluation, interpretation, and reporting of results, 899 identification from bacteria colonies, 895– 898 advanced identification, 897–898 basic identification, 895, 897 flow scheme, 896 in Acanthamoeba, 2389 isolation procedures, 892–895 microscopy, 890–891 molecular diagnosis Legionnaires’ disease, 892 specimen collection, transport, and handling, 298 taxonomy, 887 typing systems, 898 Legionella quateirensis, 888 Legionella quinlivanii, 888 Legionella rowbothamii, 888 Legionella rubrilucens, 888 Legionella sainthelensi, 888, 894 Legionella santicrucis, 888 Legionella shakespearei, 888 Legionella spiritensis, 888 Legionella steelei, 888 Legionella steigerwaltii, 888 Legionella taurinensis, 888 Legionella tucsonensis, 888 Legionella tunisiensis, 888 Legionella wadsworthii, 888 Legionella waltersii, 888 Legionella worsleiensis, 888 Legionella yabuuchiae, 888 Legionellaceae, 887

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 99

Legionellales (order), 1150 Legionnaires’ disease, 887, 890–892 antibody determination, 898 antimicrobial susceptibilities and susceptibility testing, 898–899 clinical significance, 890 epidemiology, transmission, and pathogenesis, 889 evaluation, interpretation, and reporting of results, 899 laboratory tests suggested for, 125 molecular diagnosis, 892 Leifsonia antimicrobial susceptibility testing, 1328 chemotaxonomic features, 475 description of genus, 478 identification, 438, 495 Leifsonia aquatica, 484, 495 Leishmania, 2357–2362 animal inoculation, 2361 antiparasitic agent resistance, 2554–2555 antiparasitic agent susceptibility testing methods, 2564, 2566–2567 arthropod vector, 2507 blood specimens, 2306–2307 bone marrow aspirate, 2306 clinical significance, 2358–2359 collection of specimens, 2359–2361 commercial kits for immunodetection of serum antibodies, 2296 culture, 2361 detection, 2327–2329, 2332–2335 diagnosis, 2359 direct examination, 2361 epidemiology and transmission, 2357–2358 life cycle and morphology, 2357–2358 media for culture, 2315–2316 PCR detection, 2361 prevention, 2361–2362 serologic tests, 2361 skin testing, 2361 stains for detection, 2312–2313 storage methods, 166 treatment, 2361–2362, 2530, 2542 Leishmania aethiopica, 2358, 2360 Leishmania amazonensis, 2360 Leishmania braziliensis, 2358, 2360–2361 Leishmania chagasi, 2307, 2360–2361 Leishmania colombiense, 2360 Leishmania donovani, 2307, 2358–2360 antiparasitic agent resistance, 2554–2555 detection, 2328, 2330, 2335–2336 Leishmania garnhami, 2360 Leishmania guyanensis, 2360 Leishmania infantum, 2307, 2360–2361, 2554 Leishmania killicki, 2360 Leishmania lainsoni, 2360 Leishmania major, 2358, 2360 Leishmania mexicana, 2360 Leishmania naffi, 2360 Leishmania panamensis, 2360 Leishmania peruviana, 2360 Leishmania pifanoi, 2360 Leishmania shawii, 2360 Leishmania tropica, 2358, 2360, 2554 Leishmania venezuelensis, 2360 Leishmaniasis, 2357–2362 antiparasitic agent resistance mechanisms, 2554–2555 antiparasitic agent susceptibility testing methods, 2564, 2566–2567 arthropod vectors, 2507 commercial kits for immunodetection of serum antibodies, 2296

n xcix

culture, 2307 cutaneous, 2358–2359, 2361 detection, 2332 mucocutaneous, 2358–2359, 2361 post-kala-azar dermal, 2358–2362 visceral, 2358–2362 Leiurus, 2520 Lelliottia, 723 Lelliottia amnigena, 716, 719, 723 Lelliottia nimipressuralis, 716, 719, 723 Lemierre’s disease, 905, 973 Leminorella, 715, 721, 727 Leminorella grimontii, 718 Leminorella richardii, 718 Lemniscomys virus, 1669, 1671 Lentivirus (genus), 1399, 1436, 1458 Lepidoptera (order), 2518–2519, 2522 Leprosy (Hansen’s disease), 540–541, 1360 Leptomyxid encephalitis, 2391 Leptonema, 1028 Leptonema illini, 1030 Leptosphaeria senegalensis, 2174 Leptosphaeria tompkinsii, 2174 Leptospira, 1028–1033 antibiotic susceptibilities, 1033 clinical significance, 996–997, 1030–1031 collection, transport, and storage of specimens, 283, 1031 dark-field microscopy, 9 description, 1028 direct examination, 1031 Ellinghausen-McCullough-Johnson-Harris medium for, 336 epidemiology and transmission, 1028–1030 evaluation, interpretation, and reporting of results, 1033 genotypic classification, 1028–1030 identification, 994, 997, 1031–1032 isolation procedures, 1031 microscopy, 1031 nucleic acid detection, 1031 scanning electron micrograph, 1030 serologic classification, 1028–1029 serologic tests, 1032–1033 taxonomy, 1028–1030 typing systems, 1032 Leptospira alexanderi, 1029 Leptospira alstonii, 1029 Leptospira biflexa, 997, 1028–1029 Leptospira borgpetersenii, 1029 Leptospira broomii, 1029 Leptospira fainei, 1029 Leptospira idonii, 1029 Leptospira inadai, 1029 Leptospira interrogans, 304–305, 997, 1028– 1029 Leptospira kirschneri, 1029 Leptospira kmetyi, 1029 Leptospira licerasiae, 1029 Leptospira meyeri, 1029 Leptospira noguchii, 1029 Leptospira santarosai, 1029 Leptospira terpstrae, 1029 Leptospira vanthielii, 1029 Leptospira weilii, 1029 Leptospira wolbachii, 1029 Leptospira wolffii, 1029 Leptospira yanagawae, 1029 Leptospiraceae (family), 1030, 1037 Leptospirosis, laboratory tests suggested for, 125 Leptotrichia characteristics of genus, 970–971 clinical significance, 497, 973–974

c

n

SUBJECT INDEX

Leptotrichia (continued) identification, 976–977, 979–981 taxonomy, 968 Leptotrichia amnionii, 968, 974–975, 979 Leptotrichia buccalis, 656–657, 659, 968, 974, 977, 979, 981 Leptotrichia goodfellowii, 968, 974, 979, 981 Leptotrichia hofstadii, 968, 977, 979, 981 Leptotrichia hongkongensis, 968, 974, 981 Leptotrichia shahii, 968, 977, 979, 981 Leptotrichia trevisanii, 968, 974, 977, 979, 981 Leptotrichia wadei, 968, 974, 979, 981 Leptotrichiaceae (family), 652–653, 967–968 Leptotrombidium, 2507 Leptotrombidium akamushi, 2511 Leptotrombidium deliense, 2511 Letermovir cytomegalovirus, 1720 herpesviruses, 1886 Lethargy, in tick paralysis, 2516 Leuconostoc antimicrobial susceptibilities, 430, 1184, 1189–1190 antimicrobial susceptibility testing, 1254, 1317, 1330 clinical significance, 424 epidemiology and transmission, 423 identification, 426–427, 429 interpretation of results, 431 isolation procedures, 425 taxonomy, 422 Leuconostoc citreum, 422 Leuconostoc lactis, 422 Leuconostoc mesenteroides, 422 Leuconostoc pseudomesenteroides, 422 Leucovorin calcium, for Toxoplasma gondii, 2381–2382 Leukocyte detection, 275 Leukoencephalomalacia, 2190 Leukopenia carbapenems, 1177 cephalosporins, 1175 chloramphenicol, 1193 linezolid, 1191 nitrofurantoin, 1196 sulfonamides, 1192 vancomycin, 1189 Levofloxacin, 1178–1180, 1199 antimicrobial susceptibility testing, 1256, 1260 for Mycobacteria, 1361 Lewia infectoria, 2155 Liaison HSV-1 and HSV-2 IgG tests, 1693 Liaison treponema screen, 1071–1072 Liat influenza A/B assay, 1477 Libraries for molecular epidemiology, 150– 151 strain catalogues, 150–151 surveillance databases, 151 Library subtyping, definition, 132 Lice, 2510–2511, 2522 detection, 2328, 2330 treatment, 2534 Lichtheimia, 1937, 2088, 2091–2094 Lichtheimiaceae (family), 2088, 2091–2092 Lichtheimia corymbifera, 2088, 2091–2092, 2095 Lichtheimia ornata, 2088, 2092 Lichtheimia ramosa, 2088, 2092 Lidocaine, 2520 Ligamenvirales (order), 1402 Light Diagnostics CMV pp65 antigenemia, 1724 Light Diagnostics HMPV reagent, 1503, 1511

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 100

Light Diagnostics respiratory viral panel DFA, 1503 Light Diagnostics SimulFluor HSV 1/2, 1692 Light Diagnostics SimulFluor HSV1 and HSV2 DFA typing kit, 1692 Light Diagnostics SimulFluor HSV/VZV, 1692 Light Diagnostics SimulFluor viral diagnostic screen, 1473 LightCycler, 74 herpes simplex virus (HSV), 1691 molecular detection of antibacterial resistance, 1379, 1381–1382 real-time RT-PCR for norovirus, 1626 LightCycler CMV quantitative kit, 1726 LightCycler Corynebacterium, 488 LightCycler MRSA advanced test, 361 LightCycler SeptiFast test, 407 LightCycler SeptiFast test M, 1997 LightCycler VRE assay, 407 LightCycler VZV Qual kit, 1707 Light-emitting diode (LED) microscopes, 550–551 Lim broth, 338 Lincomycin, 1185, 1199 Lincomycin resistance, 1231 Lincosamide(s), 1185 adverse effects, 1185 mechanism of action, 1185 pharmacology, 1185 spectrum of activity, 1185 Lincosamide resistance, 1231 Lindane, 2517 Lindnera jadinii, 1985, 2000 Line immunoassays (LIAs), for HTLVs, 1462–1463, 1465 Line probe assays antiviral susceptibility testing, 1916, 1918 for Mycobacterium, 581–583 Linear array, 1791, 1795 Linezolid, 1190–1191 adverse effects, 1191 antimicrobial susceptibility testing, 1256, 1261 concentration in serum, 1199 for Mycobacterium infection, 1361 mechanism of action, 1190 pharmacology, 1190 spectrum of activity, 1190–1191 Linezolid resistance, 1230–1231 in enterococci and staphylococci, 1278 molecular detection, 1385 Streptococcus, 1320 Linguatula serrata, 2516 Linnaean system of classification, 2285 Lipase, Clostridium identification, 953–954 Liponyssoides, 2507 Liponyssoides sanguineus, 2511 Lipopeptide(s), 1187–1189 adverse effects, 1189 antimicrobial susceptibility testing, 1260 mechanism of action, 1187–1188 pharmacology, 1188 spectrum of activity, 1188–1189 Lipopeptide resistance, 1188 Lipopolysaccharide (LPS) Brucella, 863–864 Coxiella burnetii, 1150 Escherichia coli, 690, 695 Pseudomonas, 780 Liquid sterilization, 205 Liquid-phase hybridization protection assay, 54 Listeria, 462–467

antimicrobial susceptibilities, 467, 1179, 1184, 1188–1189 clinical significance, 463 commercial sources of chromogenic agar media for, 326 description, 462 direct examination, 463–464 epidemiology and transmission, 462–463 evaluation, interpretation, and reporting of results, 467 identification, 438, 464–466 isolation procedures, 464 serologic tests, 467 specimen collection, transport, and storage, 463 taxonomy, 462 typing systems, 466–467 Listeria fleischmannii, 462 Listeria fleischmannii subsp. coloradonensis, 462 Listeria grayi, 462, 465–466 Listeria innocua, 462, 464–466 Listeria ivanovii, 462, 464–466 Listeria ivanovii subsp. ivanovii, 462, 465 Listeria ivanovii subsp. londoniensis, 462, 465 Listeria marthii, 462 Listeria monocytogenes, 462–467 antimicrobial susceptibilities, 467, 1172, 1176, 1182, 1188, 1190, 1192 antimicrobial susceptibility testing, 1317, 1330 Biosynth chromogenic medium for, 328– 329 clinical significance, 463 description, 462 direct examination, 463–464 epidemiology and transmission, 462–463 evaluation, interpretation, and reporting of results, 467 identification, 464–466 in Acanthamoeba, 2389 isolation procedures, 464 multistate outbreak (2011), 128 serologic tests, 467 specimen collection, transport, and storage, 463 subtyping, 139 taxonomy, 462 typing systems, 466–467 UVM (University of Vermont) modified Listeria enrichment broth for, 346 Listeria monocytogenes confirmatory base agar, 338 Listeria Oxford medium base with antibiotic inhibitor, 338 Listeria rocourtiae, 462 Listeria seeligeri, 462, 464–466 Listeria transport enrichment medium, 338 Listeria weihenstephanensis, 462 Listeria welshimeri, 462, 465–466 Listeriosis, 462–463 Lisuride, 2190 Litostomatea (class), 2287, 2416 Littman Oxgall agar, 1952, 1961, 2139 Live attenuated vaccines, 147–148 Liver, trematodes of, 2481, 2484, 2487–2490 Liver abscess/infection Campylobacter, 1001 Echinococcus granulosus, 2476–2477 Echinococcus multilocularis, 2477 Edwardsiella, 721 Entamoeba histolytica, 2331, 2403 Klebsiella pneumoniae, 718, 723 Lactococcus, 424 liver trematodes, 2489

SUBJECT INDEX Liver infusion agar, 338 Liver specimen/biopsy hepatitis B virus, 1853 hepatitis C virus, 1602–1603 parasitology, 2294, 2299, 2328, 2330–2331 Ljungan virus, 1541 Lleida bat virus, 1633–1634 Loa loa, 2461–2464, 2467–2468 arthropod vector, 2507 clinical significance, 2467 description of agent, 2467 detection, 2328, 2330 diagnosis, 2467 direct examination, 2467 epidemiology and transmission, 2467 microscopy, 2467 nucleic acid detection, 2467 prevention, 2467–2468 serologic tests, 2467 taxonomy, 2467 treatment, 2467–2468, 2531–2532, 2534– 2535 Lobosea (class), 2387 Locus for enterocyte effacement (LEE) pathogenicity island, Escherichia coli, 686, 688–689, 694–695 Loeffler medium, 338 Loiasis, 2330, 2467, 2507 Lombard-Dowell agar, 339 Lombard-Dowell egg yolk agar, 339 Lomefloxacin, 1179, 1256, 1260 Lone Star ticks, 2516 Lonomia achelous, 2518 Loop-mediated amplification (LAMP), 66, 68, 575 arboviruses, 1652 coronaviruses, 1571 liver trematodes, 2489 noroviruses, 1626–1627 Plasmodium, 2347–2348 respiratory syncytial virus, 1502 Lopinavir resistance, 1897–1898 Louse-borne relapsing fever, 1037–1041, 2511; see also Borrelia Louse-borne typhus, 1125 Lowenstein-Gruft medium, 339 Lowenstein-Jensen medium, 339, 1362 Lower respiratory tract infection, see also Pneumonia; Pulmonary disease/ infection adenoviruses, 1771 bocavirus, 1823 coronaviruses, 1569 human metapneumovirus, 1509 parainfluenza virus, 1488 respiratory syncytial virus (RSV), 1499– 1500 rhinoviruses, 1553 Lower respiratory tract specimen collection, transport, and storage guidelines, 278 fungi, 1946–1948, 1950 specimen collection, transport, and handling, 271, 297–299 bronchoscopy, 298 Chlamydia and Chlamydophila, 299 cystic fibrosis patients, 299 Legionella, 298 M. pneumoniae, 298–299 Nocardia, 299 Loxosceles, 2520 Loxosceles reclusa, 2521 Loxoscelism, 2520–2521 LPS, see Lipopolysaccharide

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 101

Luciferase immunoprecipitation system assay, for Loa loa, 2467 Lucilia sericata, 2517–2518 Lugol’s iodine, 2313 Lujo virus, 1669–1671, 1674–1676, 1679, 1681 Lumefantrine, 2539, 2564 adverse effects, 2539 mechanism of action, 2539 pharmacokinetics, 2539 spectrum of activity, 2539 Lumefantrine resistance, 2552 Luminex microbead hybridization technology Aspergillus, 2042 Fusarium, 2068 hyaline fungi, 2076–2077 Luminex xMAP flow cytometer, 59–60 Luna virus, 1669, 1671 Lung abscess Anaerococcus, 911 Ceriporia lacerata, 2075 Coprinus cinereus, 2075 etiologies, usual, 290 Gemella, 424 Lung disease of prematurity, Mycoplasma, 1092 Lung disease/infection, see also Pulmonary disease/infection Aspergillus, 2033, 2036–2037, 2044 Balantidium coli, 2417 Coprinus cinereus, 2075 Echinococcus granulosus, 2476 Mycobacterium, 542–544, 599, 1369–1370; see also Tuberculosis Mycoplasma, 1092 Paracoccidioides brasiliensis, 2115 Perenniporia, 2075 Pseudomonas, 776 Pseudomonas aeruginosa, 775 Talaromyces marneffei, 2046 Lung specimen/biopsy Mycobacterium, 547 parasitology, 2294, 2329, 2331 Pneumocystis, 2020 Luteipulveratus, 354, 361 Lutzomyia, 2507 LYI-S-2 medium, 2315 Lyme disease (Lyme borreliosis), 1037–1049; see also Borrelia arthropod vectors, 2507, 2512, 2521, 2523 specimen management, 283 LYMErix, 1048 Lymph node specimen parasitology, 2328, 2331–2332 specimen handling, 291–292 Lymphadenitis/lymphadenopathy adenoviruses, 1772 Aerococcus, 424 Bartonella, 876–877 centipede bites, 2520 Corynebacterium diphtheriae, 480 Corynebacterium pseudotuberculosis, 479 Epstein-Barr virus, 1739 herpes B virus, 1697 human herpesvirus 6 (HHV-6), 1755– 1756 human herpesvirus 8 (HHV-8), 1762– 1763 Leishmania, 2359 Mansonella, 2468 monkeypox virus, 1830–1831 Mycobacterium avium complex, 541 Mycobacterium genavense, 542 Mycobacterium haemophilum, 542

n ci

Mycobacterium kansasii, 542 Mycobacterium malmoense, 542 Mycobacterium simiae, 543 non-spore-forming, anaerobic, Grampositive rods, 923 Onchocerca volvulus, 2466 Orientia, 1124 Rickettsia, 1124 Rosai-Dorfman disease, 1756 rubella, 1526 Talaromyces marneffei, 2046 Toxoplasma gondii, 2375, 2379 Trypanosoma brucei, 2366 Trypanosoma cruzi, 2362 Yersinia pseudotuberculosis, 742 Lymphangitis filarial nematodes, 2464 herpes B virus, 1697 Lymphatic filarial nematodes, 2461–2465 antigen detection, 2465 clinical significance, 2462, 2464 description of agents, 2461 diagnosis, 2464 direct examination, 2464–2465 epidemiology and transmission, 2461–2462 Knott’s concentration technique, 2465 microscopy, 2464–2465 nucleic acid detection, 2465 nucleopore filtration, 2465 serologic tests, 2465 taxonomy, 2461 treatment and prevention, 2465 Lymphatic filariasis, 2462, 2464–2465 Lymphatics specimens, for parasitology, 2328, 2331–2332 Lymphedema, lymphatic filariasis and, 2462, 2464–2465 Lymphocryptovirus (genus), 1398 Lymphocutaneous infection, Nocardia brasiliensis and, 516 Lymphocytic choriomeningitis virus, 1669, 1671, 1673, 1675–1676, 1678–1681 Lymphocytic pleocytosis, HSV, 1689 Lymphocytoma, borrelial, 1041 Lymphocytosis, Epstein-Barr virus, 1739 Lymphogranuloma venereum, 1108, 1111, 1113, 1116 Lymphoid disorders, viral specimens, 1406 Lymphoma adult T-cell leukemia/lymphoma (ATLL), 1460–1462 Helicobacter, 1017 primary effusion lymphoma (PEL), 1763– 1764 Lymphopenia, coronavirus, 1569 Lymphotrophic polyomavirus (species), 1803 Lyophilization, see Freeze-drying (lyophilization) Lysergic acid, 2190 Lysinibacillus description of genus, 441 taxonomy, 441 Lysinibacillus sphaericus, 441, 443, 449, 452 Lysis-centrifugation blood culture system, 20, 1948–1949 Lysostaphin, 354 Lysozyme test, 319 Lyssavirus (genus) disinfectant susceptibility, 1635 diversity, 1634 epidemiology and transmission, 1633, 1635 morphology and structural proteins, 1634

cii

n

SUBJECT INDEX

Lyssavirus (genus) (continued) rabies virus, 1633–1641 taxonomic classification, 1398 Lytta vesicatoria, 2521 M2 channel blocker resistance, 1903–1904 M2 protein inhibitors, influenza virus, 1886– 1887 M-11 medium, 2315 M48, 1542 M2000 system, 74, 1110–1111 M2000rt system, 74 M2000sp system, 74 Macacine herpesvirus 1, see Herpes B virus MacConkey agar, 339 MacConkey agar, Fluorocult, 339 MacConkey agar with sorbitol, 339 Machupo virus, 1669–1671, 1673–1675, 1678, 1680–1681 Macracanthorhynchus hirudinaceus, 2291 Macrobilharzia, 2480 Macrococcus, 354–357, 361 Macrococcus caseolyticus, 356, 357, 364, 368 Macroconidia, 1941, 2060–2061, 2067–2068, 2129–2134, 2145 Macrodilution method, see Broth macrodilution method Macrolide(s), 1182–1183 adverse effects, 1183 antimicrobial susceptibility testing, 1255, 1260 fidaxomicin, 1183 for Mycobacterium infection, 1361 mechanism of action, 1182, 1361 pharmacology, 1182 spectrum of activity, 1182–1183 Macrolide resistance, 1182, 1215, 1231 Brevibacterium, 1328 Dermabacter, 1328 Haemophilus influenzae, 1321 Mycobacterium avium complex, 1357, 1361 Streptococcus, 1320 Streptococcus pneumoniae, 1316 Turicella, 1328 Macrolide-lincosamide-streptogramin B (MLSB) class of antibiotics, resistance to, 1215, 1218, 1231, 1316 Macrophage activation syndrome, adenoviruses and, 1773 Madura foot, 513 Madurella, 2174, 2176, 2181 Madurella fahalii, 2176 Madurella grisea, 2174, 2176 Madurella mycetomatis, 1968, 2161, 2174– 2182 Madurella pseudomycetomatis, 2176 Madurella tropicana, 2176 Maggots, 2518 Magicplex RV panel real-time test, 1574 MagNA Pure system, 74, 1542, 1691 Magnusiomyces capitatus, 1984, 1986, 2000 Maintenance medium, 161, 1430 Major facilitator superfamily (MFS) transporters, 2241 Malachite green broth, 339 Malacoplakia, Rhodococcus equi and, 519 Malaise adenoviruses, 1772 arboviruses, 1647 arenaviruses, 1674 coronaviruses, 1569 Corynebacterium diphtheriae, 480 Cystoisospora belli, 2428 cytomegalovirus, 1719

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 102

dirofilariasis, 2499 Ehrlichia chaffeensis, 1138 influenza virus, 1471 malaria, 2339 Rickettsia, 1124 varicella-zoster virus, 1704 Malaria, 2338–2349; see also Plasmodium antimalarials, 2536–2541 antifolates, 2540–2541 artemisinin derivatives, 2539–2540 atovaquone-proguanil, 2541 quinoline derivatives, 2536–2539 antiparasitic agent resistance mechanisms, 2550–2553 antiparasitic agent susceptibility testing methods, 2563–2566 arthropod vectors, 2507 detection, 2334–2335 diagnostic direct methods, 2347 immunochromatographic tests, 2335–2336 laboratory tests suggested for, 125 Malarone, 2349, 2541 Malassezia, 1937, 1998, 2001 antifungal susceptibilities, 2004–2005 antifungal susceptibility testing, 2263 clinical significance, 1994 description of agents, 1990 direct examination, 1995 media, 1961 media for culture, 1952 microscopy, 1966, 1969, 1973 phenotypic characteristics, 1990 specimen collection, transport, and processing, 1948–1949, 1952 taxonomy, 1985 Malassezia caprae, 1990 Malassezia cuniculi, 1990 Malassezia dermatis, 1990, 2005, 2146 Malassezia equina, 1990 Malassezia furfur, 1985, 1990, 2005, 2146– 2147 clinical significance, 1994 specimen collection, transport, and processing, 1949 Malassezia globosa, 1961, 1985, 1990, 1994, 2005, 2146 Malassezia japonica, 1990, 2005 Malassezia nana, 1990, 2005 Malassezia obtusa, 1961, 1990, 2005, 2146 Malassezia pachydermatis, 1985, 1998, 2146 antifungal susceptibilities, 2005 antifungal susceptibility testing, 2263 clinical significance, 1994 description, 1990 phenotypic characteristics, 1990 Malassezia restricta, 1961, 1990, 1994, 2005, 2146 Malassezia slooffiae, 1990, 1994, 2005, 2146 Malassezia sympodialis, 1990, 1994, 2005, 2146 Malassezia yamatoensis, 1990, 2005, 2146 Malawi polyomavirus (MWPyV), 1803, 1805, 1810 Malbranchea, 2118 MALDI Biotyper, see Biotyper system MALDI-TOF MS, 72 Achromobacter, 843 aerobic catalase-negative, Gram-positive cocci, 429 anaerobic bacteria, 906 anaerobic Gram-negative rods, 982 antifungal susceptibility testing in yeasts, 2267 Arcobacter, 1006

Aspergillus, 2042–2043 Bacillus, 454 Bacillus anthracis, 450 Bartonella, 880 Bordetella, 843 Brucella, 867 Burkholderia, 799–800 Campylobacter, 1006 Candida, 1978 carbapenemase detection, 1301 Clostridium, 954–955 Corynebacterium, 484 Enterobacteriaceae, 722 Enterococcus, 411 Escherichia, 693 for identification of bacteria, 262–263 for rapid microbial identification of blood culture isolates, 24–25 for subtyping, 145 Francisella, 858 fungal identification, 1978 Fusarium, 2068 Gram-negative nonfermentative rods, 816–817 Gram-positive anaerobic cocci (GPAC), 913 HACEK group, 658, 660–661, 676–677 Haemophilus, 676–677 Helicobacter, 1020–1021, 1024 human papillomavirus (HPV), 1795 hyaline fungi, 2077 identification of Gram-positive cocci, 352 Legionella pneumophila, 897–898 Listeria, 466 molecular detection of antibacterial resistance, 1380 Mycobacterium, 583–584, 604–605 Mycoplasma, 1096 Neisseria, 642 Nocardia, 525–526 non-spore-forming, Gram-positive, anaerobic rods, 929–930 organism identification, 29, 35–39, 48 Pseudomonas, 776 safety considerations, 178 Salmonella, 703 Staphylococcus, 365 Stenotrophomonas maltophilia, 801–802 Streptococcal species identification, 391, 392 turnaround time for testing, 46 Vibrionaceae, 768 yeasts, 2003 Yersinia, 746 Malignancy Epstein-Barr virus-associated, 1738–1740, 1746–1747 human immunodeficiency virus, 1439 human papillomavirus and, 1784–1786, 1788 Kaposi’s sarcoma, 1762–1763 Mallophaga (order), 2510, 2522 Malt extract (2%) agar, 1961 MAMA (mismatch amplification mutation assay), 1383 Mamastrovirus (genus), 1399, 1618 Mange demodectic, 2517 sarcoptic, 2516 Mannheimia haemolytica, 655 Mannitol salt agar, 339 Mannitol salt broth, 339 Mannitol selenite broth (selenite mannitol broth), 339

SUBJECT INDEX Mannitol-egg yolk-polymyxin agar, 339 Mannitol-lysine-crystal violet-brilliant green agar, 339 Mansonella, 2468 arthropod vector, 2507 clinical significance, 2468 description of agents, 2468 diagnosis, 2468 epidemiology and transmission, 2468 taxonomy, 2468 treatment and prevention, 2468 Mansonella ozzardi, 2462–2464, 2468 arthropod vector, 2507 treatment, 2468, 2534–2535 Mansonella perstans, 2462–2464, 2468, 2535 Mansonella streptocerca, 2462, 2464, 2468 detection, 2329, 2332, 2336 treatment, 2468, 2534–2535 Manual blood culture systems, 20 mar (multiple antibiotic resistance) operon, 1234 Maraviroc, 1871, 1877, 1920 Maraviroc resistance, 1447, 1448, 1897, 1899, 1920 Marburg virus, 1647, 1669–1681 antigen detection, 1676 clinical significance, 1674 collection, transport, and storage of specimens, 1675 description, 1670, 1672 direct examination, 1675–1677 epidemiology and transmission, 1670, 1673 evaluation and interpretation of results, 1681–1682 identification of virus, 1680 isolation procedures, 1678 nucleic acid detection, 1677 serologic diagnosis, 1680–1681 taxonomy, 1670, 1672 Marburgvirus (genus), 1398, 1670 MarDx T. pallidum Marblot, 1070, 1072 Maribavir cytomegalovirus, 1720 herpesviruses, 1886 Maribavir resistance, 1917 mariPOC test, 1503, 1511 Marrara, 2516 Marseille fever, 2513 Martin-Lewis agar, 339 Marvinbryantia, 921–922 Marvinbryantia formatexigens, 922 Mask, N95, 282 Mass spectrometry, see also MALDI-TOF MS for identification of bacteria, 262–263 Haemophilus, 676–677 identification and characterization of pathogens, 72–73 safety considerations, 178 MassArray, 1791 Masson-Fontana silver stain, 2057 Mast antibiotic resistance detection kit, 1300 Mastadenovirus (genus), 1398, 1769 Mastitis, Corynebacterium kroppenstedtii and, 479, 490 Mastoiditis Staphylococcus, 360 Stenotrophomonas maltophilia, 794 Material safety data sheets (MSDS), 175 Matrix-assisted laser desorption ionizationtime of flight mass spectrometry, see MALDI-TOF MS Maurer’s clefts, 2343–2344

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 103

MAX enteric bacterial panel, 690 MAX MRSA assay, 1381 MAX StaphSR test, 361 Mayaro virus, 1645 MB/BacT Alert 3D, 553 McBride Listeria agar, 339–340 McDade, Joseph, 889 McFarland standards, 1257, 1261, 1263, 1343–1344 MDR-TB assay, 584 MDST (molecular drug susceptibility testing), for Mycobacterium tuberculosis, 1356, 1367–1368 Measles atypical syndrome, 1520–1521 complications, 1520–1521 control programs, 147, 1520 countries with endemic spread of, 147 diagnosis, serologic, 1524–1525 endemic, 1519–1520 epidemiology and transmission, 1519–1520 in immunocompromised patients, 1521 infection during pregnancy, 1520 laboratory tests suggested for, 125 subacute sclerosing panencephalitis (SSPE), 1520 uncomplicated course, 1520 vaccine, 1519–1521 Measles inclusion-body encephalitis, 1521– 1522 Measles virus, 1519–1525 clinical significance, 1520–1521 complications, 1520–1521 uncomplicated clinical course, 1520 collection, transport, and storage of specimens, 1521–1522 cytopathic effect (CPE), 1522–1523 description of agent, 1519 detection and identification methods, 1434 direct examination, 1522 cytological examination, 1522 immunofluorescence assay, 1522–1523 nucleic acid detection, 1522 epidemiology and transmission, 1519–1520 evaluation, interpretation, and reporting of results, 1525–1526 genotypes, 1519 genotyping, 1523–1524 isolation and identification, 1522–1523 confirmation of isolation, 1523 serologic diagnosis, 1524–1525 ELISAs, 1524–1526 plaque-reduction neutralization (PRN) assay, 1524 specimen collection and handling, 1406– 1408, 1413–1414 taxonomy, 1519 Mebendazole, 2529–2532 adverse effects, 2531–2532 Ascaris lumbricoides, 2451, 2455 Capillaria philippinensis, 2497 Enterobius vermicularis, 2454–2455 Fasciolopsis buski, 2490 Giardia duodenalis, 2412 hookworm, 2455–2456 indications for, 2532 mechanism of action, 2530 Parastrongylus, 2499 pharmacokinetics, 2530 spectrum of activity, 2530–2531 Strongyloides stercoralis, 2458 Trichuris trichiura, 2455, 2459 MEC (minimum effective concentration), 2045

n ciii

mecA gene, 1222, 1249, 1267, 1278, 1291– 1292, 1294, 1380–1382 Mechanical tube length, 5–6 Mecillinam, 1172, 1259 Media blood culture, 17 fungi, 1951–1953 mycology, 1955–1956, 1959–1962 parasite culture, 2315–2316 recommendations for initial sample handling, 286–287 Medical devices classification, 197–198 critical items, 197 noncritical items, 197, 198 semicritical items, 197 disinfection of, 197–201 classification of devices, 197–198 dental equipment, 200–201 endoscopes, 198–200 reuse of single-use devices, 205–206 samples for fungi, 1946, 1950 Medical laboratory assistants, 46 Medical laboratory technologists (MLT), 45, 46 Medical technologists (MT), 45, 46 Medical waste, 174 Medicopsis romeroi, 1968, 2174–2175, 2178– 2182 Mediterranean spotted fever, 2513 mefA gene, 1316 Mefloquine, 2538, 2564 adverse effects, 2538 JC polyomavirus, 1811 mechanism of action, 2538 pharmacokinetics, 2538 Plasmodium, 2349 spectrum of activity, 2538 Mefloquine resistance, 2552, 2564 Megaesophagus/megacolon, Trypanosoma cruzi, 2363 Megamonas, 969 Megamonas funiformis, 969 Megamonas hypermegale, 969 Megamyxovirus (genus), 1487 Megasphaera, 497 description of, 909 epidemiology, 910 identification, 913, 916 taxonomy, 909 Megasphaera elsdenii, 909 Megasphaera micronuciformis, 909 Meglumine antimoniate, 2362, 2542, 2554, 2564 Meglumine antimoniate resistance, 2554 Meiosis, 1936 Melanconiales, 1939 Melanized fungi, 2153–2168 antifungal susceptibilities, 2167 antigen and nucleic acid detection, 2164 clinical significance, 2161–2164 collection, transport, and storage of specimens, 2164 direct examination, 2164 epidemiology and transmission, 2161 evaluation, interpretation, and reporting of results, 2167–2168 identification, 2164–2165 isolation, 2164 microscopy, 2164 serologic tests, 2167 taxonomy and description of agents, 2153–2161 typing systems, 165–2167

civ

n

SUBJECT INDEX

Melarsoprol, 2544, 2551, 2555, 2564, 2567 adverse effects, 2544 mechanism of action, 2544 pharmacokinetics, 2544 spectrum of activity, 2544 Trypanosoma brucei, 2367 Melarsoprol resistance, 2567 Melioidosis, 794–795 biothreat agent, 222 laboratory-acquired infections, 176–177 transmission and disease, 222 Membrane filtration for parasites, 2307, 2336 Menekre virus, 1669, 1671 Meningitis Achromobacter, 841 Actinobacillus, 654 adenovirus, 1778 Aeromonas, 754 arboviruses, 1647 Bacillus cereus, 443 Bacillus circulans, 443 Balneatrix alpica, 827 Bergeyella zoohelcum, 827 Blastomyces dermatitidis, 2114 Borrelia, 1041 Burkholderia, 794 Campylobacter, 1000 Candida, 1993 Capnocytophaga, 654 Chryseobacterium indologenes, 828 Citrobacter, 720 Clostridium tertium, 948 Coccidioides, 2114 Comamonas testosteroni, 795 Cronobacter sakazakii, 719 Cryptococcus, 1993 Edwardsiella, 721 Ehrlichia chaffeensis, 1138 Elizabethkingia meningoseptica, 813, 828, 831 enterovirus, 1539–1541 Escherichia coli, 686, 688 etiologies, usual, 290 Finegoldia magna, 911 Fusobacterium, 973 Gemella, 424 Globicatella, 425 Gram-negative nonfermentative rods, 813 Gram-positive anaerobic cocci (GPAC), 910–911 Haemophilus influenzae, 669 Helicobacter, 1017 herpes simplex virus (HSV), 1689 Histoplasma capsulatum, 2114 human herpesvirus 6 (HHV-6), 1755 Leptospira, 1030 Listeria monocytogenes, 463 Lysinibacillus sphaericus, 443 MIC value reporting, 1254 Mycobacterium scrofulaceum, 544 Mycobacterium tuberculosis, 538 Mycoplasma, 1092 Neisseria lactamica, 645 Neisseria meningitidis, 637, 1323–1324 Paenibacillus alvei, 443 Parastrongylus, 2498 parechovirus, 1539, 1541 Pasteurella, 655 phaeohyphomycoses, 2161, 2163 Plesiomonas shigelloides, 721 Pseudomonas, 776 Pseudomonas aeruginosa, 775 Ralstonia, 795 Rhodotorula, 1994

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 104

Serratia, 720 specimen collection, transport, and handling, 294 Sporobolomyces, 1994 Staphylococcus, 360 Stenotrophomonas maltophilia, 794 Streptococcus gallolyticus subsp. pasteurianus, 387 Streptococcus pneumoniae, 386, 1250, 1315 Streptococcus salivarius group, 387 Tsukamurella paurometabola, 519 varicella-zoster virus, 1709 Veillonella, 911 Vibrio cincinnatiensis, 766 viruses, specimens, and methods for detection of, 1407 Meningococcal meningitis, laboratory tests suggested for, 125 Meningococcal shock syndrome, 637 Meningoencephalitis adenoviruses, 1773 amebic, 2330, 2389–2390 Anaplasma phagocyrophilum, 1139 arenaviruses, 1673 Ehrlichia chaffeensis, 1138 enterovirus, 1540 Epstein-Barr virus, 1739 herpes simplex virus (HSV), 1689 mumps virus, 1493 Mycoplasma, 1091 Naegleria fowleri, 2389–2390 Parastrongylus, 2498 primary amebic meningoencephalitis (PAM), 2387, 2389–2390, 2392, 2395 rubella, 1526 Trypanosoma brucei, 2366 Trypanosoma cruzi, 2363 Mercia Syphilis M, 1067 Mercia Syphilis Total, 1068 Mercury, in parasitology preservatives/ fixatives, 2310–2312 Mercury vapor (HBO) lamps, 10 Mericitabine resistance, 1902 Meridian viral transport, 1410 Merifluor Cryptosporidium/Giardia, 2295, 2441 Merino Walk virus, 1669, 1671 Meristematic, 1941 Meristracraceae (family), 2087 Merkel cell polyomavirus (MCPyV), 242, 1803, 1805–1807, 1810 Meropenem, 1176–1177, 1199 anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1255, 1260 Bacteroides fragilis group susceptibility percentages, 1350 Merosporangium, 1941 MERS-CoV, 1565–1578 biosafety, 1570, 1577 clinical significance, 1569 collection, transport, and storage of specimens, 1570 description of agent, 1565–1566 direct detection, 1570–1577 discovery, 1567 epidemiology and transmission, 1569 evaluation, interpretation, and reporting of results, 1578 isolation procedures, 1577 nucleic acid detection, 1577 origin, 1565–1566

phylogenetic relationships, 1566 serologic tests, 1577 structure, 1565–1566 taxonomy, 1565–1566 Merthiolate-iodine-formalin, 2310–2311 Mesocestoidae (family), 2291 Mesocestoides, 2502 Mesoplasma, 1089–1090 Mesostigmata (order), 2511 Metabolome, 227 Metagenome, 227 Metagonimiasis, 2482 Metagonimus, 2482 Metagonimus yokogawai, 2320, 2449, 2490 Metallo-β-lactamase, 1176, 1178, 1226, 1300–1301, 1346 Metals, disinfection, 195 Metamonada (phylum), 2287, 2408–2409 Metapneumovirus, see Human metapneumovirus Metapneumovirus (genus), 1398, 1487, 1508 Metarhizium, 2064, 2073, 2076 Metarhizium anisopliae, 2064, 2073, 2076 Metastrongylidae (family), 2289 Metastrongyloidea (superfamily), 2289 Metatranscriptome, 227 Methenamine silver stain, 1958 Methicillin, 1255 Methicillin-resistant S. aureus (MRSA), 150 ccrB typing tool, 150 commercial sources of chromogenic agar media for, 326 decolonization, 188–189 in skin microbiome, 232 media for detection, 324 nasal and nasopharynx carriage rate, 231 rapid detection, 111 reporting to infection prevention program, 112 strain libraries, 150 subtyping, 139 surveillance cultures, 113–114 triclosan for, 186 Methyl red-VP medium, 340 Methylene blue dye test, Toxoplasma gondii, 2376, 2379–2381 Methylene blue stain, 323 Methylobacteriaceae (family), 829 Methylobacterium, 617 collection, transport, and storage of specimens, 814 identification, 827, 829–830 Methylobacterium mesophilicum, 829 Methylobacterium zatmanii, 829 4-Methylumbelliferyl-β-D-glucuronide (MUG), 318 Metorchis albidus, 2481 Metorchis conjunctus, 2481, 2531 Metronidazole, 1194 adverse effects, 1194 anaerobic bacterial susceptibility percentages, 1351 Bacteroides fragilis group susceptibility percentages, 1350 Balantidium coli, 2417 Blastocystis hominis, 2407 concentration in serum, 1199 Dientamoeba fragilis, 2413 Dracunculus medinensis, 2497 Entamoeba histolytica, 2405 Gardnerella vaginalis, 498 Giardia duodenalis, 2412 mechanism of action, 1194 pharmacology, 1194

SUBJECT INDEX spectrum of activity, 1194 Trichomonas vaginalis, 2415 trichomoniasis, 2564, 2566 Metronidazole resistance, 1231–1232 Actinomyces, 1352 Bacteroides fragilis group, 1347 Blastocystis hominis, 2407 Gram-positive, non-spore-forming rods, 1348 Gram-positive cocci, 1349, 1351 Helicobacter pylori, 1329 Prevotella, 1347 Trichomonas vaginalis, 2415, 2551, 2553– 2554 trichomoniasis, 2564, 2566 Metula, 1941 Meyerozyma, 1985, 1988 Meyerozyma caribbica, 1985 Meyerozyma guilliermondii, 1985–1986, 2000 M’Fadyean stain, 323 M’Fadyean test, 448–449 MFC (minimum fungicidal concentration), 2256–2257, 2273 MGB Alert BK virus primers and probe, 1810 MGB Alert CMV 3.0 primers and probes, 1726 MGB Alert hMPV, 1506 MGB Alert HMPV detection reagent ASR, 1511 MGB Alert Influenza A&B/RSV, 1506 MGIT 960 system, 1365–1366 MGIT SIRE kit, 1365–1366 MHA-TP test, 1066, 1070, 1074 MHT (modified Hodge test), 1225 MIBE (measles inclusion body encephalitis), 1521–1522 M.I.C. Evaluator anaerobic bacteria, 907, 984, 1345 described, 1263 MIC (minimum inhibitory concentration) antifungals, 2256 Aspergillus, 2044–2045 breakpoints, 1248–1249, 1253, 1259–1261, 1268 MIC methods, 1247–1250; see also Agar dilution susceptibility testing; Broth microdilution antimicrobial susceptibility testing antifungal susceptibility testing, 2256 breakpoints, 1248–1249, 1253, 1259–1261, 1263, 1268 distributions, 1248, 1253–1254 factors influencing measurements, 1248 fastidious bacteria, 1315, 1317–1318 interpretive categories, 1248–1249, 1253, 1259–1261, 1263, 1343–1344 pharmacological target values, 1248 potential agents of bioterrorism, 1316 quality control, 1263–1264 reporting of results, 1250 selection of antibacterial agents for routine testing, 1249–1250, 1254 zone diameters compared with, 1249 Micafungin, 2228 antifungal susceptibility testing, 2255– 2273 eumycotic mycetoma fungi, 2181–2182 hyaline fungi, 2077 melanized fungi, 2167 phaeohyphomycosis, 2167 scedosporiosis, 2167 spectrum of activity, 2224 Talaromyces marneffei, 2048

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 105

Micafungin resistance, 2239 Michaelis-Gutmann bodies, 519 Miconazole, for Naegleria fowleri, 2395 Microarrays, see also DNA microarrays; Hybridization arrays Candida, 1997, 2003 pathogen discovery, 241–42 Microascaceae (family), 2071 Microascales (order), 1937, 1938, 2153, 2155, 2159, 2162–2163, 2173–2174 Microascus, 2062, 2069, 2070, 2071, 2075 Microascus cinereus, 2062, 2075 Microascus cirrosus, 2062, 2075 Microascus manginii, 2062 Microascus trigonosporus, 2062, 2075 Microbacterium antimicrobial susceptibility testing, 1328 chemotaxonomic features, 475 clinical significance, 479 description of genus, 477–478 epidemiology and transmission, 479 identification, 438, 484, 495 Microbacterium foliorum, 495 Microbacterium oxydans, 495 Microbacterium paraoxydans, 495 Microbacterium resistens, 495–496 Microbial genomics for pathogen discovery, 240–246 microarray-based approaches, 241–242 recent history of, 240–241 sequencing-based approaches, 242–246 Microbiome, see also Human microbiome definition, 227 DNA-based studies, 76 Micrococcaceae (family), 474, 477 antimicrobial susceptibilities, 371 clinical significance, 361 description of family, 356 direct examination, 361 epidemiology and transmission, 357 evaluation, interpretation, and reporting of results, 372 identification, 366–367 isolation procedures, 362 taxonomy, 354 Micrococcus, 354–356, 361 Micrococcus halobia, see Nesterenkonia halobia Micrococcus luteus, 357, 366–367 Micrococcus lylae, 366–367 Micrococcus roseus, see Kocuria rosea Microconidia, 1941, 2060–2061, 2067–2068, 2129–2134, 2145 Microdilution antimicrobial susceptibility testing, broth, see Broth microdilution antimicrobial susceptibility testing Microfilariae, 2499 detection, 2312, 2316, 2326, 2332, 2334– 2336 filarial nematodes, 2461–2468 stains for, 2312, 2316 Microflex system, 25 non-spore-forming, anaerobic, Grampositive rods, 930 Staphylococcus, 365 Micrographica (Hooke), 5 Microhemagglutination assay for antibodies to T. pallidum (MHA-TP) test, 1066, 1070, 1074 Microhematocrit centrifugation, 2336 Microimmunofluorescence (MIF) test, for Chlamydiaceae, 1114–1116 Micrometry, 11 Micromonospora, 1181, 1220

n cv

MicroRNAs (miRNAs), 1747 Microsatellite typing Aspergillus, 2043 Talaromyces marneffei, 2046–2047 yeasts, 2004 MicroScan Streptococcus, 1320 Streptococcus pneumoniae, 1319 MicroScan MICroSTREP, 1319 MicroScan WalkAway system, 32–33, 364, 428, 1276–1277 Bartonella, 880 data management system, 1275, 1277 Enterococcus, 411 Gram-negative bacteria, 1278–1279 Gram-positive bacteria, 1277–1278 Neg ID type 2 and type 3, 768 Prompt Inoculation System, 1277 Pseudomonas, 780 rapid anaerobe panel, 880 RENOK device, 33, 1275, 1277 Synergies plus panel, 364, 1276 Microscope, see also specific microscope types anatomy of, 7 care and use of, 12 compound, 7–9 epifluorescence, 10 simple, 7 Microscopic agglutination test (MAT), for Leptospira, 1028, 1032–1033 Microscopic observation of drug susceptibility (MODS), 1356, 1367 Microscopy, see also specific microscope types dark-field, 9 ergonomics, 12–13 fluorescence, 10–11 fungal detection and identification, 1965– 1970, 1973–1976 linear measurements (micrometry), 11 phase-contrast, 9 photomicroscopy, 11–12 technical background and definition of terms, 5–6 MicroSeq 500 16S rRNA bacterial sequencing kit, 260 MicroSeq analysis software, 76 MicroSeq D2 system, 2003 MicroSeq fungal identification, 76 MicroSeq microbial identification system, 580, 602 Microsporidia, 1938–1939, 2209–2216 antimicrobial susceptibilities, 2216 chemofluorescent staining, 2214 chromotrope-based staining, 2214 clinical significance, 2213 collection, transport, and storage, 2213 description of genera and species, 2209– 2212 detection, 2325–2326, 2328–2332 detection procedures, 2214–2215 biopsy specimens, 2215 corneal scrapings, 2215 cytological diagnosis, 2214–2215 examination of stool specimens, 2214 molecular techniques, 2215 epidemiology and transmission, 2212–2213 evaluation, interpretation, and reporting of results, 2216 identification, 2215–2216 immunofluorescent-antibody tests, 2214 isolation, 2215 life cycle, 2211 serologic tests, 2216 taxonomy, 2209

cvi

n

SUBJECT INDEX

Microsporidiosis, 2213–2214, 2331 Microsporidium, 2328, 2330 Microsporidium africanum, 2210, 2212 Microsporidium ceylonensis, 2210, 2212 Microsporum, 1937, 1939, 2128–2146 anthropophilic species, 2135 antimicrobial susceptibilities, 2145 characteristics, 2129–2130 clinical significance, 2135–2136 colony characteristics, 2140, 2144 description of etiologic agents, 2145 epidemiology and transmission, 2135 evaluation, interpretation, and reporting of laboratory results, 2146 geophilic species, 2135 growth on BCPMSG, 2141 identification, 2139–2141 in vitro hair perforation test, 2140 isolation, 2138–2139 laboratory testing of specimens, 2137– 2139 media, 1961 microscopic morphology, 2140 microscopy, 2137–2138, 2140, 2143 molecular identification techniques, 2141 nucleic acid detection, 2139 nutritional requirements, 2140 physiological tests, 2140–2141 specimen collection, transport, and processing, 1944, 1947, 1953, 2136–2137 taxonomy, 2128 temperature tolerance and temperature enhancement, 2141 urea hydrolysis, 2141 zoophilic species, 2135 Microsporum audouinii, 2129, 2135–2136, 2138, 2140–2141, 2143 Microsporum canis, 1961, 2128–2129, 2135– 2136, 2138, 2140, 2143–2144 Microsporum cookei complex, 2129, 2146 Microsporum duboisii, 2130 Microsporum ferrugineum, 2129, 2135–2136, 2138, 2141, 2145 Microsporum gallinae, 2128, 2130, 2135 Microsporum gypseum complex, 2130, 2135, 2138, 2143, 2145 Microsporum mirabile, 2129 Microsporum nanum, 2130, 2135 Microsporum persicolor, 2130, 2135, 2141, 2143, 2145 Microsporum praecox, 2130, 2135, 2138 Microsporum racemosum, 2130, 2135 Microsporum vanbreuseghemii, 2130, 2135 MicroTest M4, 1410 MicroTest M4RT, 1410 MicroTest M5, 1410 MicroTest M6, 1410 MicroTrak HSV 1 & 2 culture identification/typing test, 1692 Middle East respiratory syndrome (MERS), 1565–1578 Middlebrook 7H9 broth, 1364–1367 Middlebrook 7H9 broth with Middlebrook ADC enrichment, 340 Middlebrook 7H10 agar, 1362–1366 Middlebrook 7H10 broth with Middlebrook ADC enrichment, 340 Middlebrook 7H11 broth with Middlebrook ADC enrichment, 340 Middlebrook 7H11 medium, 1366 Middlebrook albumin-dextrose-catalase (ADC) enrichment, 340 Middlebrook OADC enrichment, 340

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 106

Midichloria mitochondrii, 1136 “Midichloriaceae, Candidatus,” family, 1135 Migratory integumomyiasis, 2518 Milker’s nodules, 1829, 1831, 1837 Millipedes, 2520, 2522 Miltefosine, 2542–2543 Acanthamoeba, 2395 adverse effects, 2543 Balamuthia mandrillaris, 2395 Leishmania, 2361–2362, 2564, 2566 mechanism of action, 2542 Naegleria fowleri, 2395 pharmacokinetics, 2542 spectrum of activity, 2542–2543 Miltefosine resistance, 2551 Mineral oil, storage of microorganisms in, 162 Minocycline, 1185–1187, 1199, 1256, 1260 mip gene, Legionella pneumophila, 898 MIRU-VNTR typing, 584–585 Miscarriage, parvovirus B19 and, 1820 Mismatch amplification mutation assay (MAMA), 1383 Mites Demodex folliculorum, 2517 dust mites, 2517 pyroglyphid, 2517 scabies, 2516–2517 vectors, 2507, 2511–2512 Mitosporic, 1941 MK7655, 1178 MLMT, see Multilocus microsatellite typing (MLMT) analysis MLSA, see Multilocus sequence analysis MLST, see Multilocus sequence typing MLVA, see Multilocus variable-number tandem-repeat analysis MLVAbank, 150 MLVANET, 150–151 MMR (measles-mumps-rubella) vaccine, 1492–1493, 1521 Mobala virus, 1669, 1671 Mobillivirus (genus), 1398 Mobiluncus antimicrobial susceptibilities, 931 clinical significance, 925 identification, 926 taxonomy and description, 920–921 Mobiluncus curtisii, 920, 925–926, 931 Mobiluncus mulieris, 920, 926 Model Performance Evaluation Program, CDC, 1365 Modified acid-fast staining, 2319 Modified Columbia agar, 2315 Modified Dixon’s medium, 1953 Modified Elek method, 488 Modified Field’s stain, 2316 Modified Hodge test, 1225, 1279, 1300–1301 Modified Kinyoun stain, 2313 Modified polyvinyl alcohol, for stool specimen preservation, 2303–2304 Modified safranin stain, 2316 Modified trichrome stain, 2319 Modified Ziehl-Neelsen stain, 2313 Moellerella, 715, 721, 726 Moellerella wisconsensis, 718, 721 Mogibacterium, 921–922, 927, 930 Mogibacterium timidum, 924, 930 Mogibacterium vescum, 924 Mokola virus, 1633–1634 Molds, 1941 antifungal susceptibility testing agar dilution method, 2272 broth macrodilution method, 2269, 2270

broth microdilution method, 2268–2271 clinical breakpoints, 2270–2271 colorimetric methods, 2271–2272 disk diffusion method, 2272 Etest, 2272 gradient strip testing, 2272 molecular methods, 2272–2273 Neo-Sensitabs, 2272 classification and identification of anamorphic molds, 1939–1940 form class Coelomycetes, 1939 form class Hyphomycetes, 1939–1940 identification of, 1940 MALDI-TOF (MS) identification of, 38 Molecular beacons, 61, 62 Molecular detection anaerobic Gram-negative rods, 975–976 of antibacterial resistance, 1379–1385 aminoglycoside, 1383 anaerobic bacteria, 1383 β-lactamases in Gram-negative bacteria, 1383–1384 beta-lactam-resistant pneumococci, 1383 ceftriaxone-resistant Neisseria gonorrhoeae, 1383 fluoroquinolones, 1383 linezolid, 1385 methicillin-resistant Staphylococcus aureus, 1380–1382 mupirocin, 1385 mycobacteria, 1385 resistance targets, 1380–1385 technology, 1379–1380 trimethoprim, 1385 vancomycin-resistant enterococci, 1381–1382 Molecular drug susceptibility testing (MDST), for Mycobacterium tuberculosis, 1356, 1367–1368 Molecular epidemiology, 131–151 applications, 145–148 molecular surveillance, 145–148 data interpretation, 149–150 diversity of organism, 149–150 epidemiological context, 150 quality of data, 149 definitions, 132 dynamics of infectious disease, 147–148 geographic spread, 147 pathogen discovery/identification, 147 pathogen evolution, 147 sustained transmission, 147 vaccination issues, 147–148 future trends, 151 libraries for, 150 strain catalogues, 150–151 strain databases, 151 overview, 131 subtyping method data interpretation, 149–150 method selection, 148–149 method validation, 149 subtyping methods, 131–145 amplified fragment length polymorphism, 137–138 DNA microarrays, 144–145 forensic microbiology, 148 k-mer analysis, 143–145 mass spectrometry, 145 non-target-specific methods, 135–137 subtyping method characteristics, 131– 135 target-specific methods, 138–141

SUBJECT INDEX whole-genome sequencing MLST and binary typing, 143 whole-genome SNP typing, 141–143 Molecular line probe assays, 1356 Molecular microbiology, 54–83 applications, 74–77 disease prognosis, 76 initial diagnosis, 74–75 nucleic acid sequencing for organism identification, 75–76 therapy, duration and response to, 76– 77 automation and instrumentation, 74 future directions of, 82–83 laboratory practice, 77–82 contamination control, 78–79 credentials, 82 quality control and assurance, 79–80 regulatory and reimbursement issues, 82 results reporting and interpretation, 80– 82 specimen collection, transport, and processing, 77–78 nonamplified nucleic acid probes, 54–55 postamplification detection and analysis, 67–73 allele-specific hybridization, 67, 69 capillary electrophoresis, 67 colorimetric microtiter plate systems, 67 gel analysis, 67 hybridization arrays, 71–72 mass spectrometry, 72–73 nucleic acid sequencing, 69–71 quantitative methods, 73 signal amplification techniques, 55–57 bDNA assays, 55–56 Cleavase-Invader technology, 56–57, 58 hybrid capture assays, 56, 57 target amplification techniques, 57–67 digital PCR, 63, 64 helicase-dependent amplification (HDA), 66–67, 69 loop-mediated amplification (LAMP), 66, 68 multiplex PCR, 58–60 nested PCR, 58 PCR, 57, 59 real-time PCR, 60–63 reverse transcriptase PCR (RT-PCR), 57–58 strand displacement amplification (SDA), 64–66 transcription-based amplification methods, 63–64, 65 Molecular serotyping, 145 Molecular surveillance of tuberculosis, 136– 137 Molecular taxonomy, 131 Molecular typing, 112–113, 1776–1777 Mollaret’s syndrome, herpes simplex virus (HSV) and, 1689 Mollicutes, 1088–1101 antimicrobial susceptibilities, 1098–1100 clinical significance, 1091–1093 collection, transport, and storage of specimens, 1093 description of, 1088–1090 direct examination, 1093–1095 epidemiology and transmission, 1090–1091 evaluation, interpretation, and reporting of results, 1100–1101 identification, 1096 isolation procedures, 1095–1096 serologic tests, 1097–1098

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 107

taxonomy, 1088–1089 typing systems, 1096–1097 Molluscipoxvirus (genus), 1398, 1828–1830, 1834 Molluscum contagiosum virus, 1828–1837 antigen detection, 1833 clinical significance, 1831 description of agents, 1828 diagnostic tests, 1832 direct detection, 1832–1835 epidemiology and transmission, 1828, 1830 identification, 1835 isolation, 1835 microscopy, 1832–1833 nucleic acid detection, 1833–1835 specimen collection and handling, 1406, 1408, 1832 taxonomy, 1828–1829 Monascus, 2189 Moniliaceous, 1941 Moniliformida (order), 2291 Moniliformidae (family), 2291 Moniliformis moniliformis, 2291 Monkey B virus, see Herpes B virus Monkeypox virus, 1402 antiviral therapy, 1831–1832 clinical significance, 1830–1831 cytopathic effect (CPE), 1836 epidemiology and transmission, 1828–1829 PCR assay, 1834–1835 serologic tests, 1836–1837 Monobactam(s), 1172, 1176 adverse effects, 1176 mechanism of action, 1176 pharmacology, 1176 spectrum of activity, 1176 structure, 1172 Monobactam resistance, 1176 Monolisa Anti-HBc, 1848 Monolisa Anti-HBs, 1847 Monolisa HBc IgM, 1848 Monolisa HBsAg 3.0, 1847 Mononegavirales (order), 1398, 1402, 1633 Mononeuritis, Epstein-Barr virus and, 1739 Mononucleosis Epstein-Barr virus, 1738–1740 herpes simplex virus (HSV), 1688 Mopeia virus, 1669, 1671 Moraxella, 615, 617–618 clinical significance, 814 collection, transport, and storage of specimens, 814 isolation procedures, 815 taxonomy, 813 Moraxella atlantae, 630–631, 820–821 Moraxella boevrei, 820 Moraxella canis, 632–633, 814, 820–821, 1182 Moraxella caprae, 820 Moraxella catarrhalis, 632–633 antibiotic resistance, 1234 antimicrobial susceptibilities, 1174–1175, 1177, 1180, 1184, 1187, 1190, 1192, 1197 antimicrobial susceptibility testing, 1317, 1330–1331 β-lactamase, 1330 β-lactamase tests, 1302 clinical significance, 813–814 epidemiology and transmission, 813 identification, 820–821 reporting of, 831 Moraxella caviae, 821

n cvii

Moraxella cuniculi, 821 Moraxella lacunata, 632–633, 813–814, 820– 821, 831 Moraxella lincolnii, 632–633, 820–821 Moraxella nonliquefaciens, 632–633, 814, 820– 821 Moraxella oblonga, 821 Moraxella osloensis, 632–633, 814, 820–821 Moraxella ovis, 821 Moraxella phenylpyruvica, see Psychrobacter phenylpyruvica Moraxellaceae (family), 813 Morbillivirus (genus), 1487, 1519 Morbus errorum, 2511 Morganella antimicrobial susceptibilities, 727–730, 1175, 1178, 1186–1187 epidemiology, transmission, and clinical significance, 720–721 identification, 724–726 Morganella morganii, 728–729, 1196 antibiotic resistance, 1226–1227 endoscope contamination outbreak, 199 Morganella morganii subsp. morganii, 717, 724, 726 Morganella morganii subsp. sibonii, 717, 724 Morganella psychrotolerans, 717 Morogoro virus, 1671 Mortality adenoviruses, 1771–1773 arenaviruses, 1673 Blastoschizomyces, 1992 cytomegalovirus, 1719 Entamoeba histolytica, 2403 enteric adenoviruses, 1622 Epstein-Barr virus, 1739–1740 hepatitis E virus, 1589, 1590 herpes simplex virus (HSV), 1689 human herpesvirus 6 (HHV-6), 1755 hyaline fungi, 2075 influenza virus, 1471 measles, 1520 monkeypox virus, 1830 parainfluenza virus, 1488 phaeohyphomycoses, 2163 Plasmodium, 2340–2341 progressive multifocal leukoencephalopathy (PML), 1804 Pythium insidiosum, 2201 rotaviruses, 1622 tick paralysis, 2516 Trypanosoma brucei, 2366 Trypanosoma cruzi, 2362, 2365 variola virus, 1830 Mortierellales (order), 2087 Morular (mulberry) cells, 2366 Mosquitoes, 2505–2506, 2522 Motavizumab, for respiratory syncytial virus, 1501 Moths, 2518–2519, 2522 Motility, 616 Mouse hepatitis virus (MHV), 1566 Mouse neutralization test, for rabies virus, 1641 Mouth abscess, anaerobic Gram-negative rods, 972 Moxifloxacin, 1178–1180, 1199 anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1256, 1260 Bacteroides fragilis group susceptibility percentages, 1350 for Mycobacteria, 1361

cviii

n

SUBJECT INDEX

Moxifloxacin resistance, 1347 MP Biomedical ELISA 3.0, 1593 MRSA, see Staphylococcus aureus, MRSA MRSA Select, 1293 MRSA/SA Elite MGB assay, 361, 1381– 1382 MRSASelect medium, 340 mtrR gene, 1322 Mucicarmine stain, 1958, 1965, 1970 Mucocutaneous infection leishmaniasis, 2358–2359, 2361 Mycoplasma, 1091 syphilis, 1059 Treponema, 1059, 1061 Mucolytic agents, 1950 Mucor, 1937, 2087–2088, 2091–2093, 2097 Mucor circinelloides, 2088, 2092–2093, 2096– 2097 Mucor circinelloides var. circinelloides, 2087 Mucor indicus, 2088, 2092–2093, 2097 Mucor irregularis, 2088, 2092, 2097 Mucor ramosissimus, 2088, 2097 Mucor velutinosus, 2088, 2097 Mucoraceae (family), 2087–2088, 2092–2094 Mucoraceous fungi, 1969 antifungal susceptibility, 2224 cycloheximide inhibition, 1955 microscopy, 1966 Mucorales (order), 1936–1937, 1951, 2087– 2098; see also Mucormycosis Mucormycosis, 2087–2098 antifungal susceptibilities, 2097 antigen detection, 2090 clinical manifestations, 2089 collection, transport, and storage of specimens, 1947, 2089–2090 direct examination and histopathology, 2090 epidemiology and transmission, 2087–2089 evaluation, interpretation, and reporting of results, 2097–2098 identification, 2091–2097 molecular, 2096–2097 phenotypic, 2091–2096 isolation, 2090 microscopy, 2090 nucleic acid detection, 2090–2091 PCR-based diagnosis in serum, 2091 taxonomy, 2087 treatment, 2089 curative, 2089 prophylaxis, 2089 typing systems, 2097 Mucoromycotina (subphylum), 1936–1937, 2087 Mucosal lesions, Histoplasma capsulatum, 2114 Mucus in respiratory specimens, 1415 Mueller tellurite medium, 340 Mueller-Hinton agar, 340, 1257, 1263, 1265–1267, 1269–1270, 1288, 1300– 1301, 1317–1318, 1324, 1329 Mueller-Hinton broth, 340, 1263–1264, 1268–1269, 1288, 1321, 1330, 1370, 1372 cation-adjusted Mueller-Hinton broth (CAMHB), 1258, 1261–1262, 1269, 1290, 1296, 1298, 1317– 1318, 1321, 1324–1325, 1327– 1328, 1330 Mueller-Hinton chocolate agar, 340 Mueller-Hinton II agar, 340 Mueller-Hinton-Fastidious (MH-F), 1269 Multicentric Castleman’s disease, 1763

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 108

Multiceps, 2329, 2332 MultiCode BK virus primers, 1810 MultiCode JC virus primers, 1810 MultiCode PLx respiratory viral panel, 1506 MultiCode-RTx assays, 62, 1691 Multidrug resistance, in Enterobacteriaceae, 715 Multidrug-resistant organisms (MDROs), 111 rapid detection, 111 reporting to infection prevention program, 112 surveillance cultures, 113–115 Multilocus enzyme electrophoresis (MLEE), Leishmania, 2361 Multilocus microsatellite typing (MLMT) analysis Aspergillus, 2043 Talaromyces marneffei, 2046–2047 Multilocus sequence analysis (MLSA), 257, 260 Enterobacteriaceae (family), 714 Nocardia, 525 nontuberculous mycobacteria, rapidly growing, 603 Multilocus sequence typing (MLST), 257 Acinetobacter, 819 Aspergillus, 2043 Bacillus anthracis, 454 Burkholderia, 799, 803 Campylobacter, 1006 Clostridium, 956–957 databases, 150 described, 139 Enterobacteriaceae, 714, 726–727 Enterococcus, 412–413 Fusarium, 2061 Helicobacter, 1021 hyaline fungi, 2059 Leishmania, 2361 Leptospira, 1032 Mycobacterium tuberculosis complex, 585 Neisseria meningitidis, 640 nontuberculous mycobacteria (NTM), slow growing, 585 Paracoccidioides brasiliensis, 2120 Pseudomonas, 780 Staphylococcus aureus, 356, 368 Streptococcus, 395–396 Talaromyces marneffei, 2047 Trichomonas vaginalis, 2414 Vibrionaceae, 769 whole-genome, 143 yeasts, 2004 Multilocus variable-number tandem-repeat (MLVR), Mycoplasma, 1097 Multilocus variable-number tandem-repeat analysis (MLVA), 140 Bacillus anthracis, 454 Brucella, 867 Enterococcus, 412–413 Escherichia coli, 695 Francisella, 859 Salmonella, 704 Staphylococcus aureus, 368 Yersinia, 747 Multiple sclerosis Chlamydia pneumoniae, 1109 human herpesvirus 6 (HHV-6), 1756 Multiplex flow immunoassays, for Treponema, 1071–1072 Multiplex HPV genotyping kit, 1791 Multiplex ligation-dependent probe amplification (MLPA), respiratory syncytial virus, 1502

Multiplex PCR, 58–60 amebae, 2393 coccidia, 2430 Dientamoeba fragilis, 2413 Giardia duodenalis, 2412 human papillomavirus (HPV), 1795 mucormycosis, 2090 Multiplex RT-PCR human metapneumovirus, 1511 influenza viruses, 1476–1478 respiratory syncytial virus, 1502, 1506 rhinoviruses, 1554–1555 Multispot HIV-1/HIV-2 rapid test, 1409, 1445, 1447 Multitrans medium, 1410 Multivirulence locus sequence typing (MVLST), 139 MUM index, 260 MUMmer software, 260 Mumps, 126, 1493 Mumps virus, 1492–1495 antigen detection, 1494 clinical significance, 1493 description of agent, 1487 detection and identification methods, 1434 diagnostic methods for detection, 1489 direct examination, 1493–1494 epidemiology and transmission, 1492–1493 evaluation, interpretation, and reporting of results, 1495 isolation and identification, 1494 laboratory techniques and control interventions used in significant outbreaks, 126 microscopy, 1493–1494 nucleic acid detection, 1494 rapid cell culture, 1426 serologic tests, 1494–1495 specimen collection and handling, 1406, 1414, 1493 taxonomy, 1487 vaccine, 1492–1493 Münster University Hospital, 150 mupA gene, 1385 Mupapillomavirus (genus), 1398, 1783 Mupirocin, 1197 Mupirocin resistance, 1235, 1297, 1385 Murdochiella, 909 Murdochiella asaccharolytica, 910 Muriform cell, 1941 Murine colonic hyperplasia, 720 Murine typhus, 1124–1126, 1129–1130, 2507 Murray Valley virus, 1645 Mus minutoides virus, 1669, 1671 Musca domestica, 2517, 2519 Muscle cramping, spider envenomation and, 2520 Muscle rigidity, spider envenomation and, 2520 Muscle specimen, for parasitology, 2294, 2300, 2329, 2332 Muscoid flies, 2513 Musculoskeletal infection Abiotrophia and Granulicatella, 424 Mycobacterium kansasii, 542 Myalgia Anaplasma phagocyrophilum, 1139 arenaviruses, 1673–1674, 1674 blackfly fever, 2515 Borrelia, 1040–1041 daptomycin, 1189 Ehrlichia chaffeensis, 1138

SUBJECT INDEX filoviruses, 1674 herpes B virus, 1697 Histoplasma capsulatum, 2114 influenza virus, 1471 Leptospira, 1030 malaria, 2339 parvovirus B19, 1819 Rickettsia, 1124 rifampin, 1195 Sarcocystis, 2429 streptogramins, 1190 Trichinella, 2495 Trypanosoma cruzi, 2362 MycAssay Aspergillus, 1979, 2039 MycAssay Pneumocystis, 1979, 2024 Myceliophthora, 2064, 2073, 2076 Myceliophthora thermophila, 2064, 2073, 2076 Mycelium, 1935, 1938, 1941 Mycetoma, 2162; see also Eumycotic mycetoma Actinomadura, 513–514 clinical features, 2176–2177 Nocardia brasiliensis, 516 Nocardia cyriacigeorgica, 516 Nocardia transvalensis, 518 Nocardia veterana, 518 Streptomyces, 519 Mycobacteria 7H11 broth with Middlebrook ADC enrichment, 340 Mycobacteriaceae (family), 536 Mycobacterium; see also Nontuberculous mycobacteria (NTM), rapidly growing; Nontuberculous mycobacteria (NTM), slowly growing antibacterial resistance, 1385 antimicrobial agents for treating, 1356– 1361 amikacin, 1358–1360 aminoglycosides, 1358–1360 amithiozone, 1360 bedaquiline, 1358, 1360 capreomycin, 1358, 1360 clofazimine, 1360 cycloserine, 1360 dapsone, 1360 ethambutol, 1358–1359 ethionamide, 1358, 1360 fluoroquinolones, 1358, 1361 isoniazid, 1357–1358 kanamycin, 1358–1360 linezolid, 1361 macrolides, 1361 PA-824, 1361 p-aminosalicylic acid (PAS), 1361 pyrazinamide, 1358–1359 quinolones, 1361 rifabutin, 1359 rifampin, 1357–1359 rifapentine, 1359 streptomycin, 1358–1360 antimicrobial susceptibilities, 1177, 1179– 1181, 1190 antimicrobial susceptibility testing, 1356– 1373 drugs used for testing, 1361 M. avium complex, 1369–1370 M. kansasii, 1370–1371 M. marinum, 1370–1371 M. tuberculosis complex, 1361–1368 nontuberculous mycobacteria, 1368– 1369 rapidly growing mycobacteria, 1371– 1372 slowly growing nontuberculous mycobacteria, 1371

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 109

chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 537–544 nontuberculous mycobacteria (NTM), 541–546 nontuberculous mycobacteria (NTM), rapidly growing, 596, 598–600 nontuberculous mycobacteria (NTM), slowly growing, 543–544, 572 collection and storage of specimens, 547– 548 blood, 548 body fluids, 548 bronchial aspirates, bronchoalveolar lavage specimens, fine needle aspirates, and lung biopsy specimens, 547 gastric lavage fluids, 548 general rules, 547 inadequate specimens, 548 nontuberculous mycobacteria (NTM), rapidly growing, 600 sputum, 547 stool specimens, 548 tissues, abscess contents, aspirated pus, and wounds, 548 urine, 548 colony morphology, 536 cross-contamination, 556 culture, 551–554 agar-based media, 552 automated, continuously monitoring systems, 553 egg-based media, 552 heme-containing medium for M. haemophilum, 552 incubation, 554 liquid media, 552–553 medium selection, 553–554 Mycobacterium growth indicator tube (MGIT), 553 reporting, 554 selective media, 552 solid media, 552 storage of positive cultures, 554 description of genus, 536–537 description of species, 537–544 Mycobacterium tuberculosis complex, 538–541 nontuberculous mycobacteria, newer species, 544–546 nontuberculous mycobacteria, rapidly growing, 595–598 nontuberculous mycobacteria, slow growing, 543–544 nontuberculous mycobacteria frequently involved in human disease, 541– 543 digestion and decontamination methods CPC method, 560 NALC-NaOH method, 558–559 optimizing, 549–550 overview, 549–550 oxalic acid method, 559–560 sodium hydroxide method, 559 sulfuric acid method, 560 Zephiran-trisodium method, 559 disinfection, 191–192, 197, 200 endoscope contamination outbreak, 199 epidemiology and transmission, 537 nontuberculous mycobacteria (NTM), slowly growing, 570, 572 evaluation, interpretation, and reporting of results, 558

n cix

nontuberculous mycobacteria (NTM), 558 nontuberculous mycobacteria (NTM), rapidly growing, 606–607 nontuberculous mycobacteria (NTM), slowly growing, 587 G+C content, 536 generation time, 537 identification, 438, 576–584 colony morphology, 577–578 genotypic for NTM species, 579–580 growth rate, 577 mycolic acid analysis, 579 niacin accumulation, 577–578 nitrate reduction, 578 nontuberculous mycobacteria (NTM), rapidly growing, 601–605 nontuberculous mycobacteria (NTM), slowly growing, 576–584 phenotypic methods, 576–579 pigmentation and photoreactivity, 577 sequence database use, 578–579 temperature, 577 immunodiagnostic tests for tuberculosis, 555–556 isolation and staining procedures, 548–554 acid-fast stain procedures, 321–322, 550–551 culture, 551–554 digestion and decontamination methods, 549–550 nontuberculous mycobacteria (NTM), rapidly growing, 601 processing specimens, 548–559 MALDI-TOF (MS) identification of, 37– 38 media for ATS medium for, 327 Lowenstein-Gruft medium for, 339 Lowenstein-Jensen medium for, 339 Middlebrook 7H9 broth with Middlebrook ADC enrichment for, 340 Middlebrook 7H10 broth with Middlebrook ADC enrichment for, 340 Middlebrook albumin-dextrose-catalase (ADC) enrichment for, 340 Middlebrook OADC enrichment, 340 Petragnani medium for, 342 Wallenstein medium for, 347 morphologic characteristics, 507 nontuberculous mycobacteria (NTM), 536 antimicrobial susceptibility testing, 1368–1369 antimicrobials for treatment of, 1357, 1361 clinical significance, 541–546 colony morphology, 577–578 contamination of specimens with, 557 description of species, 541–546 epidemiology and transmission, 537 evaluation, interpretation, and reporting of results, 558 frequently involved in human disease, 541–543 newer species, 544–546 rapidly growing, susceptibility testing, 1371-1372 slowly growing, susceptibility testing, 1371 nutritional requirements and growth, 537 processing specimens, 548–559 contaminated specimens, 559

cx

n

SUBJECT INDEX

Mycobacterium; see also Nontuberculous mycobacteria (NTM), rapidly growing; Nontuberculous mycobacteria (NTM), slowly growing (continued) sterile specimens, 548–559 quality assurance, 556–558 safety, transport, and collection of specimens, 544, 546–548 collection and storage of specimens, 547–548 laboratory safety procedures, 544, 546– 548 transportation and transfer of biological agents, 546–547 slowly growing, 570–587 antigen detection, 572, 575 antimicrobial susceptibility testing, 585– 587 characteristics of species, 571–572 clinical significance, 543–544, 572 description of species, 543–544 direct examination, 572, 575–576 epidemiology and transmission, 570, 572 evaluation, interpretation, and reporting of results, 587 identification, 576–584 genotypic for MTBC species, 581–584 genotypic for NTM species, 579–580 phenotypic methods, 576–579 sequence database use, 578–579 immunodiagnostic tests, 576 laboratory characteristics, 570–587 microscopy, 572 nucleic acid detection, 575–576 properties of species, 573–574 typing systems, 584–585 susceptibility to physical and chemical agents, 537 taxonomy, 504–505, 536–537, 595–596 vaccine, 538, 555, 576 Mycobacterium abscessus, 541, 543, 550, 557, 595–607, 1187 antimicrobial agents for treatment, 1357, 1361, 1371 quaternary ammonium compound contamination, 195 Mycobacterium abscessus subsp. abscessus, 595, 597, 599–600, 603–607 Mycobacterium abscessus subsp. bolletii, 595, 597, 603–604 Mycobacterium abscessus subsp. massiliense, 595, 597, 599–601, 603–606 Mycobacterium africanum, 537–538, 570, 578, 581, 583 Mycobacterium agri, 597, 601 Mycobacterium aichiense, 597 Mycobacterium algericum, 571, 595 Mycobacterium alvei, 597, 601 Mycobacterium aromaticivorans, 597 Mycobacterium arosiense, 542, 545, 571, 573, 582–583 Mycobacterium arupense, 544–545, 571, 574 Mycobacterium asiaticum, 543, 571, 582 Mycobacterium aubagnense, 597, 603 Mycobacterium aurum, 597 Mycobacterium austroafricanum, 597 Mycobacterium avium, 192, 537, 541, 570, 578, 581–585, 1184, 2389 Mycobacterium avium complex antimicrobial agents for treatment, 1357, 1361 antimicrobial susceptibilities, 1180, 1183, 1194, 1357, 1359

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 110

antimicrobial susceptibility testing, 1369– 1370 antimicrobial agents, 1369 indications for testing, 1369–1370 quality control, 1370 reporting results, 1370 test methods, 1370 clinical significance, 541–542, 572, 599 description of species, 541–542 detection in blood, 21 epidemiology and transmission, 537, 572 identification, 576, 581–582 in Acanthamoeba, 2389 properties of, 573 reference strains, 1370 serologic tests, 605 Mycobacterium avium subsp. avium, 541, 571, 573, 580, 585 Mycobacterium avium subsp. hominissuis, 541, 571, 580, 585 Mycobacterium avium subsp. paratuberculosis, 541, 571, 573, 585 Mycobacterium avium subsp. silvaticum, 541, 573 Mycobacterium bacteremicum, 595, 597 Mycobacterium boenickei, 597, 599 Mycobacterium bohemicum, 545, 571, 573, 587 Mycobacterium bolletii, see Mycobacterium abscessus subsp. bolletii Mycobacterium botniense, 571 Mycobacterium bouchedurhonense, 542, 545 Mycobacterium bovis, 538–539, 552, 570, 578–579, 581, 583–584, 1184, 1359 Mycobacterium bovis (BCG), 538–539, 570, 578–579, 581, 584 Mycobacterium bovis BCG vaccination, 176 Mycobacterium bovis subsp. caprae, see Mycobacterium caprae Mycobacterium branderi, 545, 571 Mycobacterium brisbanense, 597 Mycobacterium brumae, 597 Mycobacterium canariasense, 596–597 Mycobacterium canettii, 538–540, 570, 578, 581, 587 Mycobacterium caprae, 538–539, 570, 579, 581, 583 Mycobacterium celatum, 543, 571, 573, 578, 582, 587 Mycobacterium chelonae, 550, 553–554, 579, 582, 596–606 antimicrobial susceptibilities, 1180, 1183– 1184, 1187, 1192, 1359, 1361 antimicrobial susceptibility testing, 1371 Mycobacterium chelonae/M. abscessus group, 595–596, 598–599, 601, 604–605 Mycobacterium chimaera, 537, 545, 571, 573, 582–583 Mycobacterium chitae, 597 Mycobacterium chlorophenolicum, 597 Mycobacterium chubuense, 597 Mycobacterium colombiense, 541, 545, 571, 573, 583 Mycobacterium confluentis, 597 Mycobacterium conspicuum, 545, 571, 577 Mycobacterium cookii, 571 Mycobacterium cosmeticum, 596–597 Mycobacterium crocinum, 597 Mycobacterium diernhoferi, 597 Mycobacterium doricum, 545, 571 Mycobacterium duvalii, 597 Mycobacterium elephantis, 597 Mycobacterium engbaekii, 544–545, 571 Mycobacterium europaeum, 545, 570–571

Mycobacterium fallax, 597 Mycobacterium farcinogenes, 571 Mycobacterium flavescens, 597 Mycobacterium florentinum, 545, 571 Mycobacterium fluoranthenivorans, 597 Mycobacterium fortuitum, 596–597, 596–600, 602–606 antimicrobial susceptibilities, 1180, 1361 antimicrobial susceptibility testing, 1371 Mycobacterium fortuitum group, 595–596, 599, 601, 604, 606, 1187, 1191, 1371 Mycobacterium fortuitum-peregrinum complex, 602, 604 Mycobacterium fragae, 545, 571 Mycobacterium franklinii, 595, 597 Mycobacterium frederiksbergense, 597 Mycobacterium gadium, 597 Mycobacterium gastri, 542, 571, 580, 587 Mycobacterium genavense, 537, 542, 552, 571, 573, 582 Mycobacterium gilvum, 598 Mycobacterium goodii, 595, 597, 599, 601– 602, 606 Mycobacterium gordonae, 192, 537, 541, 543– 544, 550, 557, 570–573, 576, 578, 580, 582–583, 585, 587, 1371 Mycobacterium growth indicator tube (MGIT), 553, 577, 1359 Mycobacterium haemophilum, 537, 542, 548, 552–554, 571, 573, 577, 582, 586, 1361, 1371 Mycobacterium hassiacum, 598 Mycobacterium heckeshornense, 545, 571, 582 Mycobacterium heidelbergense, 545, 571, 573, 583 Mycobacterium heraklionense, 544–545, 571, 574 Mycobacterium hiberniae, 544, 571, 573 Mycobacterium hodleri, 598 Mycobacterium holsaticum, 582, 598 Mycobacterium houstonense, 596–597, 599, 602 Mycobacterium immunogenum, 597, 599, 601, 604 Mycobacterium insubricum, 598 Mycobacterium interjectum, 545, 571, 573, 582–583 Mycobacterium intermedium, 545, 571, 574, 582 Mycobacterium intracellulare, 192, 541, 570– 571, 573, 581–584, 1361 Mycobacterium iranicum, 595, 597 Mycobacterium kansasii, 537, 542, 550, 570– 572, 574, 576, 580, 582–583, 586– 587 antimicrobial agents for treatment, 1357, 1361 antimicrobial susceptibilities, 1180, 1187, 1191–1192 antimicrobial susceptibility testing, 1370– 1371 Mycobacterium komossense, 598 Mycobacterium koreense, 570–571 Mycobacterium kubicae, 545, 571 Mycobacterium kumamotonense, 544–545, 571, 574 Mycobacterium kyorinense, 545, 571 Mycobacterium lacticola, 597 Mycobacterium lacus, 545, 571 Mycobacterium lentiflavum, 537, 545, 571, 574, 580, 582, 587 Mycobacterium leprae, 536–537 characteristics of, 571 clinical significance, 540–541

SUBJECT INDEX description of species, 540–541 PCR techniques, 583 treatment, 1360 Mycobacterium lepraemurium, 541, 571 Mycobacterium litorale, 595, 598 Mycobacterium llatzerense, 598 Mycobacterium longobardum, 544–545, 570– 571 Mycobacterium madagascariense, 598 Mycobacterium mageritense, 596–598 Mycobacterium mageritense/M. wolinskyi group, 596, 598 Mycobacterium malmoense, 542–543, 570– 571, 574, 582–583 Mycobacterium mantenii, 545, 571, 583 Mycobacterium marinum, 536, 542–543, 548, 550, 553–554, 570–571, 574, 577– 578, 580, 582–583, 586–587, 1186– 1187, 1191–1192 antimicrobial agents for treatment, 1357, 1361 antimicrobial susceptibility testing, 1370– 1371 Mycobacterium marseillense, 542, 545, 571 Mycobacterium massiliense, 2389; see also Mycobacterium abscessus subsp. massiliense Mycobacterium microti, 538–540, 570, 578, 581, 583 Mycobacterium minnesotense, 570–571 Mycobacterium monacense, 596–597 Mycobacterium montefiorense, 571 Mycobacterium moriokaense, 597 Mycobacterium mucogenicum, 597, 600, 604– 606, 1371 Mycobacterium mucogenicum group, 595–596 Mycobacterium mungi, 538, 540 Mycobacterium murale, 598 Mycobacterium nebraskense, 545, 571, 580, 583 Mycobacterium neoaurum, 595, 597 Mycobacterium neworleansense, 597 Mycobacterium nonchromogenicum, 544, 571 Mycobacterium noviomagense, 571 Mycobacterium novocastrense, 597 Mycobacterium obuense, 598 Mycobacterium orygis, 538, 540 Mycobacterium pallens, 598 Mycobacterium palustre, 545, 571, 582 Mycobacterium paraffinicum, 571, 582–583 Mycobacterium parafortuitum, 598 Mycobacterium paragordonae, 571 Mycobacterium parakoreense, 570–571 Mycobacterium parascrofulaceum, 545, 571, 582–583 Mycobacterium paraseoulense, 545, 571 Mycobacterium paraterrae, 571 Mycobacterium parmense, 545, 571 Mycobacterium peregrinum, 597, 604, 1371 Mycobacterium phlei, 598 Mycobacterium phocaicum, 597, 600, 603, 605 Mycobacterium pinnipedii, 538–540, 570, 578, 581 Mycobacterium porcinum, 596–597, 599–600, 602 Mycobacterium poriferae, 598 Mycobacterium pseudoshottsii, 571 Mycobacterium psychrotolerans, 598 Mycobacterium pulveris, 571 Mycobacterium pyrenivorans, 598 Mycobacterium rhodesiae, 598 Mycobacterium riyadhense, 545, 572, 583 Mycobacterium rufum, 598 Mycobacterium rutilum, 598

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 111

Mycobacterium salmoniphilum, 595, 598 Mycobacterium saskatchewanense, 545, 572, 582–583 Mycobacterium scrofulaceum, 544, 572, 574, 581–583, 1191–1192 Mycobacterium senegalense, 597, 602 Mycobacterium senuense, 544, 546, 572 Mycobacterium seoulense, 545, 572 Mycobacterium septicum, 597, 602 Mycobacterium setense, 597, 599 Mycobacterium sherrisii, 545, 570, 572 Mycobacterium shigaense, 545, 572 Mycobacterium shimoidei, 543–544, 572, 574, 582 Mycobacterium shinjukuense, 570, 572 Mycobacterium simiae, 537, 542–543, 570, 572, 574, 580–582, 585, 587, 1361 Mycobacterium simulans, 583 Mycobacterium smegmatis, 597–598, 601–602 Mycobacterium smegmatis group, 595–596, 598–601, 1371 Mycobacterium sphagni, 598 Mycobacterium stomatepiae, 572, 577 Mycobacterium szulgai, 537, 543, 572, 574, 582 Mycobacterium terrae complex, 191, 544, 557, 572, 574, 585, 595 Mycobacterium thermoresistibile, 598, 601 Mycobacterium tilburgii, 540, 545 Mycobacterium timonense, 542, 545, 572 Mycobacterium tokaiense, 598 Mycobacterium triplex, 545, 572, 582 Mycobacterium triviale, 544, 572 Mycobacterium tuberculosis, 536, 538–540 acid-fast stain procedures, 550–551 antigen detection, 575 antimicrobial susceptibilities, 1180–1181, 1183–1184, 1192, 1194 antimicrobial susceptibility testing, 1246, 1277 clinical significance, 538 collection and storage of specimens, 547– 548 blood, 548 body fluids, 548 bronchial aspirates, bronchoalveolar lavage specimens, fine needle aspirates, and lung biopsy specimens, 547 gastric lavage fluids, 548 general rules, 547 inadequate specimens, 548 sputum, 547 stool specimens, 548 tissues, abscess contents, aspirated pus, and wounds, 548 urine, 548 colony morphology, 538–539 cross-contamination, 556 culture, 551–554 agar-based media, 552 automated, continuously monitoring systems, 553 egg-based media, 552 incubation, 554 liquid media, 552–553 medium selection, 553–554 Mycobacterium growth indicator tube (MGIT), 553 reporting, 554 selective media, 552 solid media, 552 storage of positive cultures, 554 description of species, 538

n cxi

detection in blood, 21 disinfection, 191–192, 197, 200 endoscope contamination outbreak, 199 epidemiology and transmission, 537 evaluation, interpretation, and reporting of results, 558 genome sequence, 241 identification, 576–579 immunodiagnostic tests for tuberculosis, 555–556 in CSF specimen, 94 isolation and staining procedures, 548–554 acid-fast stain procedures, 550–551 culture, 551–554 digestion and decontamination methods, 549–550 processing specimens, 548–559 laboratory-acquired infections, 177 Middlebrook 7H11 broth with Middlebrook ADC enrichment for, 340 outbreak genotyping, 245 Pfizer TB medium base with glycerol, egg yolk, glucose, and malachite green for, 342 quality assurance, 556–558 rpoB gene, 580 safety, transport, and collection of specimens, 544, 546–548 collection and storage of specimens, 547–548 laboratory safety procedures, 544, 546– 548 transportation and transfer of biological agents, 546–547 subtyping of strains, 136–137 susceptibility to physical and chemical agents, 537 Mycobacterium tuberculosis complex, 536 amikacin resistance, 1356, 1360 antibiotic resistance, 1356–1368 acquired, 1357 genes associated with, 1357–1361 primary, 1357 reporting, 1363 antibiotic resistance genes, 1357–1358 antigen detection, 572, 575 antimicrobial agents for treating, 1356– 1361 antimicrobial susceptibility testing, 586– 587, 1361–1368 agar proportion method, 1363–1365 critical concentrations, 1362 drug resistance, 1361–1362 extent of service, 1362 “fall-and-rise” phenomenon, 1362 GeneXpert MTB/RIF assay, 1368 HAIN assays, 1368 low versus high critical concentrations, 1362 methods, 1362–1368 MGIT 960 system, 1365–1366 microscopic observation of drug susceptibility (MODS), 1356, 1367 molecular drug susceptibility testing, 1356, 1367–1368 PCR, 1367–1368 reporting resistance, 1363 “special-populations” hypothesis, 1362 test concentrations, 1363 TREK Sensititre MYCOTB MIC plate method, 1367 VersaTREK, 1366–1367

cxii

n

SUBJECT INDEX

Mycobacterium tuberculosis complex (continued) when to perform testing, 1362–1363 capreomycin resistance, 1356 clinical significance, 538–541, 572 culture, 552–554 description of species, 538–541 direct examination, 572, 575–576 epidemiology and transmission, 570, 572 ethionamide resistance, 1358, 1360, 1367 evaluation, interpretation, and reporting of results, 587 fluoroquinolone resistance, 361, 1357– 1358 genotypic identification, 581–584 AccuProbe test, 581–582 gyrB gene sequencing, 581 line probe assays, 581–583 mass spectrometry, 583–584 multiplex real-time PCR assays, 583 RD/spoligotyping, 581 real-time PCR assays, 581 identification, 576–584 genotypic, 581–584 phenotypic methods, 576–579 immunodiagnostic tests, 576 isoniazid resistance, 1356–1358, 1363, 1367 kanamycin resistance, 1356, 1360 laboratory characteristics, 570–587 novel proposed species within, 540–541 nucleic acid detection, 575–576 pyrazinamide resistance, 1357–1359, 1366, 1367 quinolone resistance, 361, 1357–1358, 1367 reference strain, 1365–1366 rifabutin/rifapentine resistance, 1359 rifampin resistance, 1356–1358, 1363, 1367–1368 streptomycin resistance, 1360 typing systems, 584–585 IS6110 RFLP typing, 584 MIRU-VNTR typing, 584–585 multilocus sequence typing (MLST), 585 spoligotyping, 584 Mycobacterium tuberculosis database, 151 Mycobacterium tuberculosis subsp. caprae, see Mycobacterium caprae Mycobacterium tusciae, 545, 572 Mycobacterium ulcerans, 536, 548, 570, 587 characteristics of, 572 clinical significance, 543 culture, 552–554 decontamination protocols, 549 description of species, 543 generation time, 537 genotypic identification, 580 identification, 582–583 properties of, 574 Mycobacterium vaccae, 598 Mycobacterium vanbaalenii, 598 Mycobacterium vulneris, 541, 545, 572–573 Mycobacterium wolinskyi, 596–599, 601, 606 Mycobacterium xenopi, 199, 537, 543, 570, 572, 574, 577, 580, 582–583, 1180, 1187 Mycobacterium yongonense, 545, 570, 572 Mycobactosel agar, 340 Mycobactosel L-J medium, 340 Mycobiotic agar, 1952, 1961, 2139 Mycocladiaceae (family), 2091 Mycocladus, 2091

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 112

Mycocladus corymbifera, 2088 MYCOFAST Evolution2, Evolution3, and RevolutioN, 1099 Mycofast US, 1096 Mycolic acid analysis, Mycobacterium, 579 Mycolic acids, 1357, 1360 Myco-Lit-ATB, 1099 Mycology, see Fungi; specific fungal agents Mycoplasma, 1088–1101 antimicrobial susceptibilities, 1098–1100, 1184 methods used, 1098–1099 susceptibility profiles and treatment, 1099–1100 clinical significance, 1083, 1091–1093 genitourinary infections, 1091–1092 neonatal infections, 1092 respiratory infections, 1091 systemic infections and immunosuppressed hosts, 1092– 1093 collection, transport, and storage of specimens, 1093 specimen type and collection, 1093 transport and storage, 1093 description of, 1088–1090 diagnostic tests, 1085–1086 direct examination, 1093–1095 antigen detection, 1094 microscopy, 1093–1094 nucleic acid detection, 1094–1095 epidemiology and transmission, 1083, 1090–1091 evaluation, interpretation, and reporting of results, 1100–1101 identification, 1096 isolation procedures, 1095–1096 biosafety considerations, 1095 commercial media and culture kits, 1096 development of colonies, 1096 growth media and inoculation, 1095 incubation conditions and subcultures, 1095–1096 PPLO agar for, 342 serologic tests, 1085–1086, 1097–1098 taxonomy, 1088–1089 typing systems, 1096–1097 Mycoplasma agar base (PPLO) agar base, 340 Mycoplasma amphoriforme, 1088–1089, 1091 Mycoplasma broth base without crystal violet and with ascitic fluid, 340–341 Mycoplasma buccale, 1089 Mycoplasma faucium, 1089 Mycoplasma fermentans antimicrobial susceptibilities, 1099 clinical significance, 1091, 1093 colonization, metabolism, and pathogenicity, 1089 epidemiology and transmission, 1090 isolation, 1096 Mycoplasma genitalium antigen detection, 1094 antimicrobial susceptibilities, 1099 clinical significance, 1083, 1091–1093 collection, transport, and storage of specimens, 1093 colonization, metabolism, and pathogenicity, 1089 description of, 1088–1089 diagnostic tests, 1085 epidemiology and transmission, 1083, 1090 evaluation, interpretation, and reporting of results, 1101

nucleic acid detection, 1094 Mycoplasma hominis A8 agar for, 327 antigen detection, 1094 antimicrobial susceptibilities, 1099–1100, 1180, 1183, 1186–1187 biosafety considerations, 1095 clinical diseases associated with, 1083 collection, transport, and storage of specimens, 1093 colonies, 1088, 1090 colonization, metabolism, and pathogenicity, 1089 description of, 1088–1089 diagnostic tests, 1085 epidemiology and transmission, 1083, 1090–1091 evaluation, interpretation, and reporting of results, 1100 identification, 1096 isolation procedures, 1095–1096 microscopy, 1094 nucleic acid detection, 1095 serologic tests, 1085, 1098 urogenital Mycoplasma broth base for, 346 Mycoplasma IgG and IgM ELISA test system, 1098 Mycoplasma IST2, 1099 Mycoplasma lipophilum, 1089 Mycoplasma orale, 1089 Mycoplasma penetrans, 1089–1091 Mycoplasma pirum, 1089, 1091 Mycoplasma pneumoniae antigen detection, 1094 antimicrobial susceptibilities, 1099–1100, 1183, 1187, 1190 biosafety considerations, 1095 clinical significance, 1083, 1091 colonies, 1088, 1090 colonization, metabolism, and pathogenicity, 1089 description of, 1088–1089 development of colonies, 1096 diagnostic tests, 1086 epidemiology and transmission, 1083, 1090–1091 evaluation, interpretation, and reporting of results, 1100 growth media and inoculation, 1095 identification, 1096 incubation conditions and subcultures, 1095–1096 isolation procedures, 1095–1096 nucleic acid detection, 1094–1095 serologic tests, 1086, 1097–1098 specimen collection, transport, and handling, 298–299 typing systems, 1096–1097 Mycoplasma pneumoniae antibody (MP) test system, 1098 Mycoplasma pneumoniae IgG/IgM antibody test system, 1098 Mycoplasma primatum, 1089 Mycoplasma salivarium, 1089, 1093 Mycoplasma spermatophilum, 1089 Mycoplasmataceae (family), 1089 Mycoplasmatales (order), 1089 Mycoscreen Plus, 1096 Mycosel, 1947, 1949, 1952, 1961 MycoTB panel, 586 Mycotoxins, 2188–2192 bioterrorism, 2192 chemical classification and biosynthesis, 2188

SUBJECT INDEX aflatoxins, 2188–2189 citrinin, 2189 cyclopiazonic acid, 2189 ergot alkaloids, 2189–2190 fumonisins, 2189–2190 ochratoxins, 2189–2190 patulin, 2189–2190 trichothecenes, 2189–2190 zearalenone, 2189–2190 food safety, 2190–2192 biological control, 2192 climate change, effects of, 2191 common food substrates, 2191 current practices, 2192 detection, 2191 future, 2192 sick building syndrome, 2192 taxonomy of mycotoxin-producing fungi, 2191 MycoTrans specimen transport system, 2137 Mycoviruses, 2192 MycXtra fungal DNA extraction kit, for Pneumocystis, 2024 Myelitis herpes B virus, 1697 herpes simplex virus (HSV), 1689 Mygalomorphae, 2520 Myiasis, 2516–2519 accidental, 2516–2517, 2519 aural, 2518 detection, 2328, 2330 facultative, 2516, 2519 furuncular, 2517 obligate, 2516, 2519 oral or nasal, 2518 wound, 2518 Myla software, 1276 Myocarditis adenoviruses, 1773 arenaviruses, 1674 Bartonella, 874 Campylobacter, 1000 Chlamydia psittaci, 1109 Corynebacterium diphtheriae, 480 enterovirus, 1540–1541 heterophyid trematodes, 2490 human herpesvirus 6 (HHV-6), 1756 measles, 1521 microsporidia, 2210 parechovirus, 1541 specimen selection, 1541 Toxoplasma gondii, 2375 Trichinella, 2495 Trypanosoma cruzi, 2362 viruses, specimens and methods for detection of, 1406 Myonecrosis, clostridial, 945–946 C. perfringens, 945–946 C. septicum, 945 Myopathy, daptomycin and, 1189 Myopericarditis, influenza virus, 1471 Myositis influenza virus, 1471 microsporidia, 2210, 2213 Sarcocystis, 2429, 2431 Staphylococcus, 360 Myriodontium, 2064, 2073 Myriodontium keratinophilum, 2064 Myroides identification, 823–824 taxonomy, 813 Myroides odoratimimus, 626–627, 823–824 Myroides odoratus, 823–824 Myrothecium, 2190

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 113

N,N-Dimethyl-naphthylamine, 319 NAATs, see Nucleic acid amplification tests N-Acetyl-L-cysteine, 1950, 1956 NaCl agar, 341 NaCl requirement, for aerobic Gramnegative bacteria identification, 616 Nacovirus (genus), 1617 Naegleria, 887, 2387–2395 detection, 2329 stains for detection, 2312, 2316 storage methods, 166 Naegleria fowleri, 2387–2395 animal inoculation, 2394 antigen detection, 2392 clinical and laboratory diagnosis, 2392– 2393 clinical significance, 2389–2390 collection, handling, and storage of specimens, 2391–2392 culture, 2307, 2393–2394 description of agents, 2388–2389 detection, 2327, 2329 direct examination, 2392 enflagellation experiment, 2393 epidemiology, 2389 evaluation, interpretation, and reporting of results, 2395 isolation procedures, 2393–2394 media for culture, 2315–2316 microscopy, 2388, 2390 nucleic acid detection, 2392–2393 permanently stained preparations, 2392 serology, 2394 taxonomy, 2387 treatment, 2395, 2542 Nafcillin, 1171, 1199 Naftifine resistance, 2239 Nail infection Chaetomium globosum, 2075 Malassezia, 1994 Onychocola canadensis, 2076 onychomycosis, 2136–2137 Scopulariopsis brevicaulis, 2075 Nail specimen, for fungi, 1945, 1947, 1949 Nairovirus (genus), 1399, 1645, 2507 NALC-NaOH method, 320, 558–559 Nalidixic acid, 1178, 1199, 1260 Nannizziopsiaceae (family), 2073 Nannizziopsis, 2073, 2076 Nannizziopsis vriesii, 2063, 2076 Nanophyetiasis, 2482, 2490 Nanophyetus salmincola, 2482, 2488, 2490 Nanosphere Verigene, 1380–1382, 1381, 1383–1384 Naples virus, 1651 Nasal cavity specimen collection, transport, and storage guidelines, 279, 300 fungi, 1947, 1950 Gram stain and plating medium recommendations, 286 Nasal congestion, adenoviruses and, 1771 Nasal discharge, Linguatula serrata and, 2516 Nasal infection anaerobic Gram-negative rods, 972 Rhinosporidium seeberi, 2198, 2205 Nasal NK cell lymphoma, 1738 Nasal obstruction, coronaviruses and, 1569 NASBA, see Nucleic acid sequence-based amplification Nasopharyngeal carcinoma, 1738, 1741, 1746 Nasopharyngeal lymphoma, 1738 Nasopharynx specimen

n cxiii

collection, transport, and handling, 279, 300 Gram stain and plating medium recommendations, 286 parasitology, 2329, 2332 National Antimicrobial Resistance Monitoring System for Enteric Bacteria (NARMS), 705, 1327 National Center for Biotechnology Information (NCBI), 1402 National Center for Infectious Diseases, 177 National Electronic Disease Surveillance System, 121 National Healthcare Safety Network, 106 National Nosocomial Infections Surveillance (NNIS) system, 107 Nattrassia, 2146 Nattrassia mangiferae, 1951, 2057, 2153 Nausea arenaviruses, 1673 Balantidium coli, 2417 carbapenems, 1177 centipede bites, 2520 cephalosporins, 1175 clavulanic acid, 1177 Cryptosporidium, 2438 Cyclospora cayetanensis, 2428 Dientamoeba fragilis, 2413 filoviruses, 1674 hookworm, 2456 linezolid, 1191 macrolides, 1183 metronidazole, 1194 monobactams, 1176 nitrofurantoin, 1196 rifaximin, 1195 Sarcocystis, 2429 scorpion venom, 2520 spider envenomation, 2520 Strongyloides stercoralis, 2457 sulbactam, 1178 sulfonamides, 1192 telavancin, 1189 telithromycin, 1185 tetracyclines, 1187 Trichinella, 2495 NDM β-lactamases, 1383–1384 Nebovirus (genus), 1617 Necator americanus, 2323, 2454–2456 clinical significance, 2456 description, 2454 eggs, 2454 larvae, 2454 worms, 2454 diagnosis, 2456 epidemiology and prevention, 2454 taxonomy, 2454 transmission and life cycle, 2456 treatment, 2455–2456 Necrobacillosis, 973 Necrotizing enterocolitis Clostridium perfringens, 943–944 Cronobacter sakazakii, 719 noroviruses, 1622 Necrotizing fasciitis collection, transport, and storage of clinical specimens, 948 Gram-positive anaerobic cocci (GPAC), 910 Rothia mucilaginosa, 361 sample handling, 292 Staphylococcus, 360 Streptococcus pyogenes, 385 Necrotizing pneumonitis, adenovirus, 1773

cxiv

n

SUBJECT INDEX

Nectria, 1937, 2071 Nectriaceae (family), 2059 Negative control, 80 Negative predictive value, 92–93 Negativicoccus description of, 909 identification, 916 taxonomy, 909 Negativicutes (class), 968 Negri body, 1638 Neisseria, 635–646, 652 antibiotic resistance, 1232, 1234 antimicrobial susceptibilities, 642–644, 1172–1173, 1176–1177, 1181, 1186, 1193, 1197 clinical significance, 636–637 collection, transport, and storage of specimens, 637–638 description of genus, 635 direct examination, 638–640 antigen detection, 638 microscopy, 638 nucleic acid detection, 638–640 epidemiology and transmission, 635–636 evaluation, interpretation, and reporting of results, 644–646 identification, 640–642 carbohydrate utilization assays, 640–641 chromogenic enzyme substrate tests, 641 colonial morphology, 640 definitive identification, 640–641 DNA sequencing, 642 hybridization test, 642 immunologic methods, 641 MALDI-TOF MS, 642 microscopic morphology, 640 multitest identification systems, 641– 642 oxidase test, 640 presumptive identification, 640 isolation procedures, 640 Martin-Lewis agar for, 339 Mueller-Hinton agar for, 340 serologic tests, 642 taxonomy, 635 Thayer-Martin medium, 345 typing systems, 642, 643 Neisseria animalis, 636 Neisseria animaloris, 636, 641, 644, 652, 661 Neisseria bacilliformis, 635, 636, 641, 644–645 Neisseria CDC group EF-4a, 652, 661 Neisseria cinerea, 635, 636, 640, 641, 642 Neisseria denitrificans, 636 Neisseria dentiae, 636 Neisseria elongata, 635, 636, 640, 641, 645 Neisseria elongata subsp. elongata, 641 Neisseria elongata subsp. glycolytica, 641, 652, 661 Neisseria elongata subsp. nitroreducens, 635, 641, 652, 661 Neisseria flavescens, 635, 636, 640, 641, 645 Neisseria gonorrhoeae, 636, 641 antibiotic resistance, 1222, 1234, 1322– 1323, 1383 antigen detection, 638 antimicrobial resistance, 643 antimicrobial susceptibilities, 642–643, 1172, 1174–1175, 1178, 1180– 1181, 1190, 1195, 1197 antimicrobial susceptibility testing, 1265, 1322–1323 commercial test methods, 1323 incidence of resistance, 1322–1323 reference test methods, 1323

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 114

strategies for testing and reporting results, 1323 β-lactamase, 1302 β-lactamase tests, 1302 carbohydrate utilization assay, 640–641 chromogenic enzyme substrate tests, 641 clinical significance, 636–637 colonial morphology, 640 description, 635 epidemiology and transmission, 635–636 evaluation, interpretation, and reporting of results, 644 hybridization test, 638, 642 identification, 640–642 immunologic methods for culture confirmation, 641 isolation procedures, 640 MALDI-TOF-MS, 642 microscopy, 638, 640 nucleic acid amplification tests, 638–639 nucleic acid detection, 638–639 oxidase test, 640 penicillinase-producing (PPNG), 1323 reference strains, 1315 specimen collection, transport, and handling, 295–296, 300, 637 subtyping, 139 throat specimens for, 300 transport medium, Stuart for, 345 typing systems, 642 Neisseria lactamica, 636, 641, 645 Neisseria macacae, 636 Neisseria meningitidis antibiotic resistance, 1222, 1234 antigen detection, 638 antimicrobial resistance, 644, 1323–1324 antimicrobial susceptibilities, 643–644, 1172, 1180, 1183, 1190, 1193, 1195, 1197 antimicrobial susceptibility testing, 1265, 1323–1324 incidence of resistance, 1323–1324 strategies for testing and reporting results, 1324 carbohydrate utilization assay, 640–641 chromogenic enzyme substrate tests, 641 clinical significance, 637 collection, transport, and storage of specimens, 300, 637–638 colonial morphology, 640 description, 635 epidemiology and transmission, 636 evaluation, interpretation, and reporting of results, 644 identification, 640–642 immunologic methods for culture confirmation, 641 isolation procedures, 640 laboratory safety issues for handling of meningococcal cultures, 638 MALDI-TOF MS, 642 microscopy, 638, 640 nucleic acid detection, 640 oxidase test, 640 typing systems, 642, 643 vaccine, 636 Neisseria meningitidis group A, 126 Neisseria meningitidis medium, 341 Neisseria mucosa, 636, 641, 645 Neisseria PET, 641 Neisseria polysaccharea, 636, 641, 645 Neisseria shayeganii, 635 Neisseria sicca, 635, 636, 641, 645 Neisseria subflava, 635, 636, 641, 645

Neisseria subflava bv. flava, 641, 645 Neisseria subflava bv. perflava, 641, 645 Neisseria subflava bv. subflava, 641, 645 Neisseria weaveri, 635, 636, 641, 645–646, 652, 661 Neisseria zoodegmatis, 636, 641 Neisseriaceae (family), 635, 652–653 Neisseria/Haemophilus identification cards, 676, 678 Neisseriales, 635 Nelfinavir, 1871, 1876 Nelfinavir resistance, 1897–1898 Nelson’s medium, 2315 Nemathelminthes (phylum), 2448, 2495– 2496 Nematoda (class), 2289, 2461, 2465, 2467– 2468 Nematodes, 2448–2460 collection, transport, and storage of specimens, 2459 culture of larval-stage, 2321–2323 agar plate for S. stercoralis, 2322 Baermann technique, 2322 filter paper/slant culture technique, 2323 Harada-Mori filter paper strip culture, 2322–2323 detection, 2323 direct examination, 2459 egg sizes, 2449 evaluation, interpretation, and reporting of results, 2459–2460 filarial, 2461–2468 intestinal, 2448–2460 less common, 2493–2501 taxonomy and classification, 2288–2289, 2448 treatment, 2531 Nemonoxacin, 1179 Neocarpenteles, 2030 Neocosmospora, 2059 Neocosmospora vasinfecta, 2060, 2061 Neodermata (subphylum), 2471, 2473–2475 Neoehrlichia lotoris, 1136–1137 Neoehrlichia mikurensis, 1135–1139, 1144– 1145 Neofusicoccum, 2153 Neolecta, 2015 Neomycin, 1181 Neomycin blood agar, 341 Neonatal infection Acinetobacter, 813 adenoviruses, 1773 Candida, 1993 Chryseobacterium indologenes, 828 cytomegalovirus, 1718–1719 Elizabethkingia meningoseptica, 828 enteroviruses, 1538–1540 herpes simplex virus (HSV), 1688–1690, 1696 human immunodeficiency virus, 1438 Listeria monocytogenes, 463 Malassezia, 1994 Mycoplasma, 1092 parainfluenza virus, 1488 parechovirus, 1539 respiratory syncytial virus (RSV), 1500 specimen selection, 1541 Streptococcus agalactiae, 386, 388 Toxoplasma gondii, 2380–2381 Trichomonas vaginalis, 2414 viruses, specimens and methods for detection of, 1407 Neonatal sepsis

SUBJECT INDEX Bacillus idriensis, 443 Bacillus infantis, 443 Bacillus pumilus, 443 Campylobacter, 1000 parechovirus, 1539, 1541 specimen selection, 1541 Neopetromyces, 2030 Neorickettsiosis, 1139 Neorickettsia, 1135–1139 Neorickettsia helminthoeca, 1136–1138 Neorickettsia risticii, 1136–1138 Neorickettsia sennetsu, 1135–1139 Neosartorya, 1937, 2030, 2040 Neosartorya fischeri, 2031–2033 Neosartorya fumigata, 2032, 2034 Neosartorya pseudofischeri, 2031–2032, 2034, 2042 Neosartorya spinosa, 2031–2032 Neosartorya udagawae, 2031–2032, 2034, 2042 Neoscytalidium, 2146, 2153 Neoscytalidium dimidiatum, 1939, 1951, 2057, 2153–2154, 2156, 2162–2163, 2166– 2167 Neo-Sensitabs molds, 2272 yeasts, 2267 Neotestudina rosatii, 1967, 2173–2174, 2176, 2178–2180 Neotyphodium, 2190 Neozygitomycetes (class), 2087 Nephritis Campylobacter, 1000 microsporidia, 2210, 2213 penicillins, 1173 varicella-zoster virus, 1705 Yersinia pseudotuberculosis, 742 Nephropathia epidemica, 1662=1664 Nephropathy Loa loa, 2467 polyomavirus-associated nephropathy, 1804–1807, 1811–1812 Nephrotic syndrome, Plasmodium malariae, 2341 Nephrotoxicity aminoglycosides, 1181–1182 bacitracin, 1197 daptomycin, 1189 polymyxins, 1193 rifampin, 1195 sulfonamides, 1192 vancomycin, 1189 Nessler reagent, 316 Nested PCR, 58 coronaviruses, 1571 influenza viruses, 1476 varicella-zoster virus, 1710 Nesterenkonia, 354, 356, 361 Nesterenkonia halobia, 356 Neufeld test (Quellung reaction), 395 Neuraminidase (NA), influenza virus, 1470, 1481–1482 Neuraminidase inhibition assay, 1914, 1916 Neuraminidase inhibitor(s), 1887 Neuraminidase inhibitor resistance, 1903– 1905 Neuritis, Corynebacterium diphtheriae, 480 Neuroborreliosis, 1041–1043, 1047, 1049 Neurobrucellosis, 865 Neurocysticercosis, 2474, 2476, 2477 Neurologic disease/disorders arenaviruses, 1673–1674 cytomegalovirus, 1719 Epstein-Barr virus, 1739

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 115

hepatitis E virus, 1590 human herpesvirus 6 (HHV-6), 1754– 1756, 1760–1761 human herpesvirus 7 (HHV-7), 1761 influenza virus, 1471 malaria, 2339–2340 Mansonella, 2468 mucormycosis, 2089 Parastrongylus, 2498 phaeohyphomycoses, 2161, 2163–2164 Trichinella, 2495 varicella-zoster virus, 1705–1706, 1709 viruses, specimens and methods for detection of, 1407 Neuromuscular paralysis, aminoglycosides and, 1182 Neuropathy HTLV-1-associated myelopathy/tropical spastic paraparesis (HAM/TSP), 1460–1462 linezolid, 1191 nitrofurantoin, 1196 Neuroretinitis, Bartonella, 873, 877 Neurotoxicity cephalosporins, 1175 penicillins, 1173 quinolones, 1180 spider envenomation, 2520 Neurotropic arboviruses, 1647–1648 Neutralization assay, 97 adenovirus, 1776–1777 arboviruses, 1654–1655 arenaviruses, 1680 coronaviruses, 1577 filoviruses, 1680 hantaviruses, 1664 influenza viruses, 1481 mumps virus, 1495 parainfluenza virus, 1492 plaque reduction neutralization (PRN) assay for measles virus, 1524 plaque reduction neutralization (PRN) for rubella virus, 1529–1530 poxviruses, 1835–1837 rabies virus, 1641 respiratory syncytial virus, 1508 Neutropenia penicillins, 1173 sulfonamides, 1192 Neutropenic enterocolitis, Clostridium tertium, 948, 951 Nevirapine, for human immunodeficiency virus (HIV), 1870, 1874 Nevirapine resistance, 1897–1898 New York City medium, 341 New York virus, 1661 Next-generation sequencing (NGS), 70–71, 141 Human Microbiome Project, use of, 226 pathogen discovery, 241–247 NGU (nonchlamydial, nongonococcal urethritis), 1091, 1098, 1100 Niacin accumulation, Mycobacterium and, 577–578 Niclosamide Diphyllobothrium latum, 2473 Dipylidium caninum, 2501 Taenia saginata, 2474 Taenia solium, 2475 Nidovirales (order), 1398, 1402, 1565 Nifurtimox, 2545 adverse effects, 2545 mechanism of action, 2545 pharmacokinetics, 2545

n cxv

spectrum of activity, 2545 Trypanosoma cruzi, 2365 Nifurtimox resistance, 2551, 2555 Niger seed base medium, 1952 Nigrograna, 2174 Nigrograna mackinnonii, 2174 Nigrosin, 1957, 1970 NIH medium, 2315 Nikkomycin Z, 2216 Nikolsky’s sign, 360 nim genes, 1347, 1383 NimbleGen arrays, 71, 144 90-60 rule, 1246 Ninhydrin reagent, 318 Nitazoxanide, 2535–2536 adverse effects, 2536 Balantidium coli, 2417 Cryptosporidium, 2442 Cyclospora cayetanensis, 2431 Cystoisospora belli, 2431 Giardia duodenalis, 2412 indications for, 2536 mechanism of action, 2535 pharmacokinetics, 2535–2536 spectrum of activity, 2536 Nitrate broth, 341 Nitrate reductase, 616 Nitrate tests, 319 Mycobacterium, 578 yeast identification, 2001 Nitrite reductase, 616 Nitrocefin, 1302–1303, 1346 Nitrofurantoin, 1195–1196, 1199, 1256, 1261 Nitrofurantoin resistance, 1232 Nitroimidazole drugs, 1361 Nits, 2510–2511 Nivalenol, 2190, 2192 Nix, 2516 NNN medium with Offutt’s modifications, 2315 Nocardia acid-fast stain, 321, 550–551 antibiotic resistance, 1234 antimicrobial susceptibilities, 1177, 1180, 1186, 1190, 1192 antimicrobial susceptibility testing, 1372– 1373 broth microdilution breakpoints, 1372 clinical significance, 1372 quality control, 1373 reporting of results, 1373 testing method, 1372–1373 chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 511, 513–518 description of genus, 506, 508 direct examination, 520–521 G+C content, 536 identification, 438, 522–527 biochemicals, 523 cell wall and cell membrane analysis, 523 gene sequencing, 524–525 genus assignment, 522 MALDI-TOF MS, 525–526 molecular, 523–526 multilocus sequence analysis (MLSA), 525 PCR with amplicon detection, 523 PCR with restriction endonuclease analysis, 523–524 pyrosequencing, 525 species assignment, 522–523

cxvi

n

SUBJECT INDEX

Nocardia (continued) susceptibility testing, 523 isolation procedures, 521–522 media, 1952 microscopy, 520–521 morphologic characteristics, 507 nucleic acid detection, 521 specimen collection, transport, and handling, 299, 520, 1948 taxonomy, 504–505 Nocardia abscessus, 512–513, 523–524 Nocardia africana, 514–518 Nocardia amikacinitolerans, 514 Nocardia anaemiae, 514 Nocardia aobensis, 514, 517 Nocardia araoensis, 514 Nocardia arthritidis, 514, 524 Nocardia asiatica, 514, 524 Nocardia asteroides, 506, 508, 511, 514–518, 526, 1191 Nocardia beijingensis, 514 Nocardia blacklockiae, 514, 518 Nocardia brasiliensis, 513, 516, 523, 527, 1372 Nocardia brevicatena, 514, 516, 521, 524 Nocardia carnea, 510, 514 Nocardia caviae, 517 Nocardia concava, 514 Nocardia corynebacterioides, 514 Nocardia cyriacigeorgica, 510, 513, 515–516, 523–524, 526–527, 1372 Nocardia elegans, 514, 517, 524 Nocardia exalbida, 514 Nocardia farcinica, 506, 508, 512–513, 515– 517, 524, 526–527, 1372 Nocardia harenae, 514 Nocardia higoensis, 514, 524 Nocardia ignorata, 514 Nocardia inohanensis, 514 Nocardia kruczakiae, 514, 517, 524 Nocardia mexicana, 514 Nocardia mikamii, 514 Nocardia neocaledoniensis, 514 Nocardia niigatensis, 514 Nocardia ninae, 514 Nocardia niwae, 514 Nocardia nova, 513, 515–518, 524, 527, 1373 Nocardia otitidiscaviarum, 510, 513, 517, 523 Nocardia paucivorans, 514, 516, 524 Nocardia pneumoniae, 514 Nocardia pseudobrasiliensis, 513, 516–517 Nocardia puris, 514 Nocardia shimofusensis, 514, 524 Nocardia sienata, 514, 524 Nocardia takedensis, 514 Nocardia terpenica, 514 Nocardia testacea, 514, 524 Nocardia thailandica, 514 Nocardia transvalensis, 514, 516, 518 Nocardia vermiculata, 514 Nocardia veterana, 512–513, 517–518, 524 Nocardia vinacea, 514 Nocardia wallacei, 513, 515, 518, 526–527, 1372 Nocardia yamanashiensis, 514 Nocardiopsis chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 518 description of genus, 508 identification, 438, 522–523 morphologic characteristics, 507 taxonomy, 505 Nocardiopsis dassonvillei, 513, 518

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 116

Nodular cutaneous lesions, microsporidia and, 2210, 2213 Noma, 654–655 Nomenclature fungi, 1936 major groups of bacteria, 258–259 name changes, 264 uncultured bacteria, 259 valid publication of bacterial names, 263– 264 Nonchlamydial, nongonococcal urethritis (NGU), 1091, 1098, 1100 Noncompetitive EIAs, 98, 99 Nonfermentative Gram-negative rods, 813– 831 antimicrobial susceptibilities, 830–831 clinical significance, 813–814 collection, transport, and storage of specimens, 814 description of agents, 813 direct examination, 814–815 epidemiology and transmission, 813 evaluation, interpretation, and reporting of results, 831 identification, 815–830 overview, 815–816 oxidase-negative GNF, 816–820 oxidase-positive, indole-negative, trypsin-negative GNF, 820–822 oxidase-positive, indole-negative, trypsin-positive GNF, 822–826 oxidase-positive, indole-positive GNF, 826–829 pink-pigmented GNF, 827, 829–830 isolation procedures, 815 taxonomy, 813 Nonfermenters, 613–634 biochemical characteristics, 624–633 dichotomous algorithms for identification, 619–623 identification of aerobic Gram-negative bacteria, 613–634 test methods, 613–616 Nonnucleoside inhibitor (NNI) resistance, HCV, 1902–1903 Nonnucleoside reverse transcriptase inhibitors (NNRTIs) for HIV, 10, 1870, 1873–1874 resistance, 1897–1898 Nonnutrient agar with live or dead bacteria, 2315 Nontuberculous mycobacteria (NTM), rapidly growing antimicrobial agents for treatment of, 1357, 1360 antimicrobial susceptibilities, 605–606 antimicrobial susceptibility testing, 1371– 1372 interpretive criteria, 1372 quality control, 1371 reporting results, 1371 test method, 1371 clinical significance, 596, 598–600 bone and joint infections, 599 central nervous system disease, 599 community-acquired skin and soft tissue infections, 596, 598 corneal infections (keratitis), 599–600 disseminated cutaneous disease, 598– 599 health care-associated infections, 600 otitis media, 600 pulmonary infections, 599 collection, transport, and storage of specimens, 600

currently recognized species, 597–598 description of species, 595–598 direct examination, 600–601 microscopy, 600–601 nucleic acid detection, 601 evaluation, interpretation, and reporting of results, 606–607 identification, 601–605 antimicrobial susceptibility tests, 601 biochemical testing, 601 carbohydrate utilization, 601 HPLC, 601 molecular methods, 601–605 isolation procedures, 601 molecular identification methods, 601–605 hsp65 gene, 602–603 MALDI-TOF MS, 604–605 multilocus sequence analysis (MLSA), 603 nucleic acid probes, 601–602 PCR-restriction enzyme analysis (PRA), 603 pyrosequencing, 604 rpoB gene, 603 sequence analysis, 602 16S rRNA gene sequence, 602 variable-number-tandem-repeat (VNTR) analysis, 603–604 serologic tests, 605 taxonomy, 595–596 typing systems, 605 enterobacterial repetitive intergenic consensus PCR (ERIC PCR), 605 pulsed-field gel electrophoresis (PFGE), 605 repetitive-sequence-based PCR (repPCR), 605 VNTR analysis, 605 Nontuberculous mycobacteria (NTM), slowly growing antimicrobial susceptibility testing, 586, 1371 characteristics of species, 571–572 clinical significance, 543–544, 572 description of species, 543–544 direct examination, 572, 575–576 epidemiology and transmission, 570, 572 evaluation, interpretation, and reporting of results, 587 genotypic identification, 579–580 complete genome sequences, 579 hsp65 gene, 580 ITS 1 region, 580 rpoB gene, 580 16S rRNA gene, 579–580 23S rRNA gene, 580 genotyping, 585 IS1245/IS900, 585 multilocus sequence typing (MLST), 585 pulsed-field gel electrophoresis, 585 repetitive-unit sequence-based PCR (rep-PCR), 585 variable-number tandem repeat (VNTR), 585 identification, 576–584 colony morphology, 577–578 genotypic for NTM species, 579–580 niacin accumulation, 577–578 phenotypic methods, 576–579 pigmentation and photoreactivity, 577 sequence database use, 578–579 laboratory characteristics, 570–587

SUBJECT INDEX nucleic acid detection, 575–576 properties of species, 573–574 Norfloxacin, 1178–1179, 1199, 1256, 1260 Norovirus (genus), 1399, 1617 Noroviruses antigen detection, 1623–1625 cell culture, 1627 clinical significance, 1620–1622 description of agents, 1619 detection and identification methods, 1434 electron microscopy, 1619, 1623 epidemiology and transmission, 1620–1621 evaluation, interpretation, and reporting of results, 1628–1629 isothermal amplification assays, 1626– 1627 molecular detection assays, 1625–1627 PCR, 1626 taxonomy, 1617–1618 typing systems, 1628 vaccine, 1622 North American deer ticks, 2512 North Asian tick typhus, 1125 Nosema, 2209–2211, 2328–2330, 2332 Nosema algerae, 2209 Nosema connori, 2209 Nosema corneum, 2210 Nosema ocularum, 2209 Nosocomial infections, see also Health careassociated infections (HAIs) arenaviruses, 1670 Asaia, 829 Aspergillus, 2031–2033 Citrobacter, 720 Elizabethkingia meningoseptica, 828 phaeohyphomycoses, 2164 Pseudomonas aeruginosa, 774–775 respiratory syncytial virus (RSV), 1500 Stenotrophomonas maltophilia, 794 Nosocomiicoccus, 354, 356–357, 361 Nosocomiicoccus ampullae, 357 Nosopsyllus, 2507 “No-touch” automated room disinfection (NTD), 196–197 Novy-MacNeal-Nicolle (NNN) medium, 2315 NS5A inhibitor resistance, HCV, 1902–1903 Nucleic acid, 69–71 chain termination methods, 69–70 CLIP sequencing, 69 high-throughput shotgun, 75 identification of bacteria and fungi by, 75–76 Mycobacterium, 602 next-generation sequencing, 70–71 ABI SOLID, 70 Ion Torrent, 70–71 Roche 454, 70 pyrosequencing, 70, 76 Nucleic acid amplification methods, for bacteremia or fungemia detection, 23, 24 Nucleic acid amplification tests (NAATs) arboviruses, 1648–1652 Borrelia, 1043 Campylobacter, 1002 Chlamydia pneumoniae, 1112 Chlamydia psittaci, 1112 Chlamydia trachomatis, 1110–1112 Chlamydiaceae, 1110–1112 Clostridium difficile, 950 coronaviruses, 1570–1578 enteroviruses, 1542–1543

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 117

Epstein-Barr virus (EBV), 1742–1743 for Streptococcus, 388–389 human metapneumovirus (HMPV), 1506– 1507 Mycobacterium tuberculosis, 557, 575 Mycobacterium tuberculosis complex, 575– 576 Neisseria gonorrhoeae, 638–639, 1323 nontuberculous mycobacteria, 576 parechoviruses, 1542–1543 polyomaviruses, 1807, 1809–1810 respiratory syncytial virus, 1502, 1506– 1508, 1511–1512 specimen handling for, 289 varicella-zoster virus, 1707–1710 Nucleic acid detection adenovirus, 1774–1775, 1778 amebae, pathogenic and opportunistic free-living, 2392–2393 Anaplasma phagocyrophilum, 1143 arenaviruses, 1677 Aspergillus, 2039–2040 Babesia, 2351–2352 Bartonella, 877 Candida albicans, 2415 coccidia, 2430 Coxiella burnetii, 1153–1154 cytomegalovirus (CMV), 1411, 1723–1726 dermatophytes, 2139 dimorphic fungi causing systemic mycoses, 2116–2117 Diphyllobothrium latum, 2473 Ehrlichia chaffeensis, 1140 Entamoeba histolytica, 2405 Enterococcus, 407 entomophthoromycosis, 2100 Epstein-Barr virus, 1739, 1741–1743 Escherichia coli, 690–691 eumycotic mycetoma, 2178 filoviruses, 1677 Francisella, 856–857 fungal identification and diagnosis, 1979 Fusarium, 2068 gastroenteritis viruses, 1625–1627 Giardia duodenalis, 2412 hantaviruses, 1663 hepatitis A virus, 1591 hepatitis B virus, 1411, 1846, 1849 hepatitis E virus, 1591–1592 human bocavirus, 1824 human metapneumovirus, 1511 hyaline fungi, 2076–2077 Hymenolepis nana, 2476 Leptospira, 1031 Loa loa, 2467 lymphatic filarial nematodes, 2465 measles virus, 1522 melanized fungi, 2164 mucormycosis, 2090–2091 mumps virus, 1494 Mycoplasma, 1094–1095 Onchocerca volvulus, 2466 parainfluenza virus, 1489–1491 parasites, 2308 parvovirus B19, 1819, 1821–1822 Plasmodium, 2347–2348 Pneumocystis, 2024–2025 polyomaviruses, 1807, 1809–1810 poxviruses, 1833–1835 Pseudomonas, 776 Pythium insidiosum, 2202 rabies virus, 1637, 1640 rhinoviruses, 1554–1555 rubella virus, 1527

n cxvii

Staphylococcus, 361–362 Taenia saginata, 2474 Taenia solium, 2475 Talaromyces marneffei, 2046 Toxoplasma gondii, 2375–2376 transmissible spongiform encephalopathies (TSEs), 1863–1864 Trichinella spiralis, 2495 Trichomonas vaginalis, 2415 Tropheryma whipplei, 1162–1163 yeasts, 1997–1998 Yersinia, 744 Nucleic acid extraction, 77–78 Nucleic acid hybridization tests, for Chlamydiaceae, 1110, 1112 Nucleic acid lateral flow immunoassay (NAL-FIA), for Plasmodium, 2348 Nucleic acid probes for Mycobacterium, 601–602 nonamplified, 54–55 Nucleic acid sequence-based amplification (NASBA), 63–64 human metapneumovirus, 1511 influenza viruses, 1476 noroviruses, 1626–1627 Plasmodium, 2348 respiratory syncytial virus, 1502, 1506 Nucleic acid sequencing, see Sequencing Nucleic acid tests hepatitis C virus, 1603–1605 herpes simplex virus (HSV), 1690–1691 human immunodeficiency virus, 1441– 1443, 1449–1451 human T-cell lymphotropic viruses (HTLVs), 1461–1462 influenza viruses, 1476–1478 specimen storage and processing, 1411– 1412 transport medium for, 1409–1410 Nucleopore filtration, for lymphatic filarial nematodes, 2465 Nucleoside inhibitor resistance, 1902–1903 Nucleoside RT inhibitors, 1440 Nucleoside/nucleotide analogues, for HBV, 1880–1882 Nucleoside/nucleotide reverse transcriptase inhibitors (NRTIs) for HIV, 1869–1870, 1872–1873 resistance in HBV, 1900 resistance in HIV, 1896–1898 Nucleotide sequence analysis, rabies virus, 1641 Nucleotidyltransferases, 1383 NucliSENS EasyQ, 64, 74, 1383–1384 Enterovirus, 1542–1543 HMPV, 1506, 1511 HPV, 1790 molecular detection of antibacterial resistance, 1379, 1381–1382 RSV A+B, 1506 Nugent scoring system, 926 Numerical aperture (NA), 6, 7 Nupapillomavirus (genus), 1398, 1783 Nutrient agar, 1.5%, HiVeg with ascitic fluid, 341 Nuttalliellidae (family), 2512 NV-AD norovirus test, 1624 Nygmia phaeorrhoea, 2518 Nystatin resistance, 2239, 2243 N-Z-amine A glycerol agar, 341 O antigen, Salmonella, 703 O157:H7 ID agar, 341

cxviii n

SUBJECT INDEX

Objectives (objective lens) labeling, 6 numerical aperture, 6, 7 types, 8–9 Obligate myiasis, 2516, 2519 Observa software, 1276 Occupational Safety and Health Administration, 282 Oceanimonas, 752 Ochratoxins, 2189–2192 Ochrobactrum, 615, 823–824 Ochrobactrum anthropi, 626–627, 823–824 Ochrobactrum haematophilum, 823–824 Ochrobactrum intermedium, 626–627, 823–824 Ochrobactrum pseudogrignonense, 823–824 Ochrobactrum pseudintermedium, 823–824 Ochroconis, 2153, 2161, 2168 Ochroconis gallopava, 2161, 2268–2269 Ocozocoautla de Espinosa virus, 1669, 1672 Octenidine, 186 Ocular fluid specimens, for Toxoplasma gondii, 2375 Ocular infection adenovirus, 1413, 1772 Alcaligenes faecalis, 841 Aspergillus, 2032 Delftia acidovorans, 795 etiologies, usual, 290 Gemella, 424 herpes simplex virus (HSV), 1689 Onchocerca volvulus, 2466 parasitic, 2330 phaeohyphomycoses, 2162 Pythium insidiosum, 2201 Shewanella, 825 Stenotrophomonas maltophilia, 794 toxoplasmosis, 2375, 2381, 2382 Vibrio alginolyticus, 765 viruses, specimens and methods for detection of, 1407 zoster ophthalmicus, 1705 Ocular objective, 9 Odontogenic infections anaerobic Gram-negative rods, 972, 974 Atopobium, 925 Fusobacterium, 973 Prevotella, 972 Odoribacter characteristics of genus, 970–971 identification, 977–978 taxonomy, 967–969 Odoribacter denticanis, 967 Odoribacter laneus, 967, 978 Odoribacter splanchnicus, 967, 978 Oerskovia antimicrobial susceptibility testing, 1328 description of genus, 477 epidemiology and transmission, 479 identification, 438, 495 taxonomy, 474–475 Oerskovia turbata, 477, 484, 495, 1328 Oerskovia xanthineolytica, 477; see also Cellulomonas cellulans Oestrus, 2330 Ofloxacin, 1178–1179, 1199 antimicrobial susceptibility testing, 1256, 1260 for Mycobacteria, 1361 Oil, storage of microorganisms in, 162, 166 Oleandomycin, 1182 Oleic acid-albumin-dextrose-catalase (OADC), 1364, 1367, 1371 Oligacanthorhynchida (order), 2291 Oligacanthorhynchidae (family), 2291

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 118

Oligella, 614 identification, 821 taxonomy, 813 Oligella ureolytica, 632–633, 820–821 Oligella urethralis, 632–633, 820–821 Oligokaryotic cell, 1941 Oliveros virus, 1669, 1671 Olsenella clinical significance, 925 enzyme reactions, 929 taxonomy and description, 920–921 Olsenella profusa, 925, 929 Olsenella uli, 925, 929 Omadacycline, 1185–1187 Ombitasvir, 1879–1880 OmniLog Phenotype MicroArray system, 778 Omsk hemorrhagic fever virus, 1645, 2507 Onchocerca volvulus, 1139, 2461–2467 arthropod vector, 2507 clinical significance, 2466 description of agent, 2466 detection, 2329, 2332 diagnosis, 2466 direct examination, 2466 epidemiology and transmission, 2466 microscopy, 2466 nucleic acid detection, 2466 prevention, 2467 serologic tests, 2466 taxonomy, 2465 treatment, 2466–2467, 2531, 2534–2535, 2544 Onchocerciasis, 2330, 2332, 2466, 2507 Onchocercidae (family), 2289, 2465, 2467– 2468, 2499 Onchocercomata, 2466 Oncosphere, 2475, 2477 OncoTect E6/E7 mRNA, 1789 ONE broth-Listeria (Oxoid novel enrichment broth-Listeria), 341 O-Nitrophenyl-β-D-galactopyranoside (ONPG), 317–318 Önöz Salmonella agar, 341 Onychocola, 2064, 2073, 2076 Onychocola canadensis, 2064, 2073, 2076 Onychomycosis, 2136–2137 Chaetomium globosum, 2075 Fusarium, 2058 Malassezia, 1994 Onychocola canadensis, 2076 Scopulariopsis brevicaulis, 2075 Onygenaceae (family), 2109 Onygenales (order), 1937, 1938, 2109, 2128, 2196 O’nyong-nyong virus, 1645 Oocysts, Toxoplasma gondii, 2373–2374 Oomycetes (class), 2200, 2203 Oomycota (phylum), 1936, 1939 Oophoritis etiologies, usual, 290 mumps virus, 1493 Oospore, 1941 OPA, 192 Operational taxonomic unit, 227 Ophiostoma, 2159 Ophiostomatales (order), 1938, 2153, 2155, 2159, 2162, 2166–2167 Ophthalmic infection Abiotrophia and Granulicatella, 424 Capnocytophaga, 654 Ophthalmomyiasis, 2518 Opinavir, for human immunodeficiency virus (HIV), 1871, 1876

Opisthokonta (supergroup), 1936 Opisthorchiasis, 2481, 2489–2490, 2533 Opisthorchiidae (family), 2290, 2481, 2487– 2488 Opisthorchioidae (superfamily), 2290 Opisthorchis, 2320, 2328, 2330 Opisthorchis felineus, 2449, 2481, 2488–2489 Opisthorchis guayaquilensis, 2481 Opisthorchis viverrini, 2481, 2488–2490, 2531 Opportunistic fungi, cycloheximide inhibition of, 1955 Optic neuropathy chloramphenicol, 1193 linezolid, 1191 Optical mapping, 136 Optical train, 7 OptiRead automated fluorometric plate reading system, 1277 Optochin test, for Streptococcus pneumoniae, 394 Oral cavity specimens fungi, 1946, 1950 specimen collection, transport, and storage guidelines, 279 viruses, 1414 Oral hairy leukoplakia, 1739 Oral human papillomavirus, 1785–1786 Oral infection Actinomyces, 922 anaerobic Gram-negative rods, 972 Campylobacter, 997 Gram-positive anaerobic cocci (GPAC), 910–911 herpes simplex virus (HSV), 1688–1689 non-spore-forming, anaerobic, Grampositive rods, 923 Parvimonas micra, 910–911, 911 Peptostreptococcus anaerobius, 911 Porphyromonas, 971 Prevotella, 972 Selenomonas, 974 Oral microbiome, 227–229 Oral mucosal transudates (OMT), 1414 measles virus, 1521 rubella virus, 1527 OraQuick Advance rapid HIV-1/2 antibody test, 1445 Orbital cellulitis, Haemophilus influenzae, 669 Orbivirus (genus), 1399 Orchitis mumps virus, 1493 viruses, specimens and methods for detection of, 1408 Orf, 1828–1831, 1835, 1837 orfX gene, 1380 Oribacterium, 921 Oribacterium sinus, 922 Orientia, 1122–1130 antimicrobial susceptibilities, 1129 clinical significance, 1083, 1124–1125 collection, transport, and storage of specimens, 1124, 1126 description of, 1123 diagnostic tests, 1086 direct detection, 1127–1128 immunologic detection, 1127 molecular detection, 1127–1128 epidemiology and transmission, 1083, 1124 identification, 1128 interpretation and reporting of results, 1130 isolation procedures, 1128 phylogeny, 1123

SUBJECT INDEX serologic tests, 1086, 1128–1129 taxonomy, 1122 Orientia chuto, 1122 Orientia tsutsugamushi, 2511 antimicrobial susceptibilities, 1129 arthropod vector, 2507 clinical significance, 1083, 1124–1125 collection, transport, and storage of specimens, 1124, 1126 description of, 1123 diagnostic tests, 1086 direct detection, 1127–1128 epidemiology and transmission, 1083, 1124 identification, 1128 immunologic detection, 1127 isolation procedures, 1128 molecular detection, 1127–1128 serologic tests, 1086, 1128–1129 taxonomy, 1122 Oritavancin, 1187–1189, 1199 Ornidazole, 1199, 2412 Ornithobilharzia, 2480 Ornithodoros, 1037–1040, 2507, 2515 Ornithodoros coriaceus, 2512 Ornithodoros erraticus, 2515 Ornithodoros hermsi, 2515 Ornithodoros moubata, 2515 Ornithodoros turicata, 2515 Oropharyngeal specimens, for fungi, 1946, 1950 Oropouche virus, 1645 Oropsylla, 2507 Oroya fever, 876 Ortho 3.0 EIA, 1608 Ortho Vitros Anti-HIV 1+2, 1444 Orthobunyavirus (genus), 1399, 1645 Orthohepadnavirus (genus), 1398 Orthomyxoviridae (family), 1470, 1644 taxonomic classification, 1398, 1400 virion morphology, 1401 Orthopolyomavirus (genus), 1803 Orthopoxvirus (genus), 1398, 1402, 1828– 1829, 1833–1835 Orthopoxviruses, 1828–1837 antigen detection, 1833 clinical significance, 1830–1832 antiviral therapy, 1831–1832 cowpox virus, 1831 monkeypox virus, 1830–1831 vaccinia virus, 1831 variola virus, 1830 collection, transport, and storage of specimens, 1832 cytopathic effect (CPE), 1835–1836 description of agents, 1828 diagnostic tests, 1832 direct detection, 1832–1835 epidemiology and transmission, 1828–1829 cowpox virus, 1829 monkeypox virus, 1829 vaccinia virus, 1829 variola virus, 1829 evaluation, interpretation, and reporting of results, 1837 identification, 1835 isolation, 1835 microscopy, 1832–1833 nucleic acid detection, 1833–1835 serologic tests, 1835–1837 specimen collection and handling, 1406 taxonomy, 1828–1829 Orthoreovirus (genus), 1399 Orthoretrovirinae (subfamily), 1399

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 119

Oseltamivir, 1471, 1481, 1886–1887, 1914– 1916 Oseltamivir resistance, 1903–1905, 1915, 1917, 1921 Osom Influenza A&B, 1474 Osom Trichomonas rapid test, 2327, 2415 Osteochondritis, Stenotrophomonas maltophilia and, 794 Osteomyelitis Aeromonas, 754 anaerobic Gram-negative rods, 972 Blastomyces dermatitidis, 2114 Blastoschizomyces, 1992 Brucella, 865 Burkholderia, 794 Capnocytophaga, 654 Chrysosporium zonatum, 2076 Coccidioides, 2114 etiologies, usual, 291 Fusarium, 2058 Gemella, 424 Histoplasma capsulatum, 2114 Inonotus tropicalis, 2075 Kingella, 655 Lactococcus, 424 Leuconostoc, 424 Myceliophthora thermophila, 2076 Mycobacterium, 542–544, 596, 599 Mycoplasma, 1093 Pantoea, 719 Parvimonas micra, 911 Pasteurella, 655 Pseudomonas, 776 Pseudomonas aeruginosa, 775 Ralstonia, 795 Salmonella, 701 Shewanella, 825 Staphylococcus, 360, 1327 Veillonella, 911 Osteoperiostitis, Treponema and, 1061 Ostiole, 1942 Otitis Delftia acidovorans, 795 etiologies, usual, 290 Otitis externa etiologies, usual, 290 Pseudomonas aeruginosa, 775 Staphylococcus, 360 Otitis media adenoviruses, 1771 Alloiococcus otitis, 361 Clostridium, 948 etiologies, usual, 290 Finegoldia magna, 911 Fusarium, 2058 Haemophilus influenzae, 669 Haemophilus parainfluenzae, 670 influenza virus, 1471 measles, 1520 Moraxella, 814 Mycobacterium, 600 Parvimonas micra, 911 rhinoviruses, 1553 Shewanella, 825 Staphylococcus, 360 Streptococcus pneumoniae, 386 Otobius, 2514 Otocentor, 2514 Ototoxicity aminoglycosides, 1182 macrolides, 1183 vancomycin, 1189 Ouchterlony analysis, 96 Outbreak

n cxix

agencies involved in response, 121 defined, 120, 132 detection, 121–123 disease surveillance, 120–121 information resources, 128–129 investigation, 120–129 epidemic curve examples, 123 epidemiological approaches, 123–124 example scenarios, 126–128 laboratory role in, 124–126 molecular surveillance and, 145–146 steps in, 123 notifiable (mandatory reportable) diseases, 121, 122 point source, 123 recognition and investigation, 112, 113 Outbreak Database, 128 Ova and parasite (O&P) examination, 2304–2305, 2317–2321 “Owl’s eye” inclusion, 1423, 1722 OXA (oxacillin hydrolyzing) β-lactamases, 1223–1224, 1227–1228.1300, 1302, 1383–1384 Oxacillin, 1171–1172, 1199 AAC-1 β-lactamase inhibition by, 1300 antimicrobial susceptibility testing, 1255, 1259 Oxacillin resistance detection by automated antimicrobial susceptibility testing, 1278 detection in Staphylococcus, 1289–1294 by PCR, 1293–1294 chromogenic agars, 1293 in coagulase-negative staphylococci, 1292–1293 in S. aureus and S. lugdunensis, 1289– 1292 rapid tests, 1293 reporting results of tests, 1294 in staphylococci, 1278, 1289–1294 Oxacillin-salt agar screening test, 1291 Oxalic acid, 320, 559–560, 1950 Oxazolidinone(s), 1190–1191, 1361 adverse effects, 1191 mechanism of action, 1190 pharmacology, 1190 spectrum of activity, 1190–1191 Oxazolidinone resistance, 1215, 1230–1231 Oxford agar, modified (Listeria selective agar, modified Oxford), 341 Oxford agar (Listeria selective agar, Oxford), 341 Oxford University, 150 Oxidase test, 319, 640 Oxidase-negative GNF, identification of, 817–820 Oxidase-positive/indole-negative/trypsinnegative GNF, identification of, 820– 822 Oxidase-positive/indole-negative/trypsinpositive GNF, identification of, 822– 826 Oxidase-positive/indole-positive GNF, identification of, 826–829 Oxidation-fermentation medium, HughLeifson’s, 341 Oxidation-fermentation medium, King’s, 341 Oxoid Brilliance Candida agar, 1960 Oxoid Salmonella chromogenic agar (OSCM), 341 Oxoid Signal system, 20 Oxyporus, 2062 Oxyporus corticola, 2062 Oxysporus corticola, 2071

cxx

n

SUBJECT INDEX

Oxyuridae (family), 2289 Oxyuroida (order), 2289 Oxyuroidea (superfamily), 2289 Ozena, 718 Ozenoxacin, 1179–1180 Ozone, low-temperature sterilization by, 205 P agar, 342 PA-824, 1361 Paecilomyces, 2069, 2073, 2076, 2077 antifungal susceptibility testing, 2268– 2269 key phenotypic features, 2064 microscopy, 1967, 1969 Paecilomyces formosus, 2064, 2073 Paecilomyces fumosoroseus, 2064 Paecilomyces fusisporus, 2073 Paecilomyces inflatus, 2071 Paecilomyces javanicus, 2064 Paecilomyces lilacinus, 2073, 2077 Paecilomyces marquandii, 2073 Paecilomyces variotii, 2064, 2073, 2076, 2077, 2261, 2270, 2272 Paederus fusca, 2521 Paenalcaligenes clinical significance, 841 collection, transport, and storage of specimens, 842 description of genus, 840 evaluation, interpretation, and reporting of results, 845 identification, 843 taxonomy, 838 Paenalcaligenes hominis, 838, 840–841, 843, 845 Paenibacillaceae, 441 Paenibacillus clinical significance, 443 description of genus, 441 epidemiology and transmission, 442 identification, 438, 451, 453 taxonomy, 441 Paenibacillus alvei, 443, 449, 451–453 Paenibacillus glucanolyticus, 443 Paenibacillus konsidensis, 443 Paenibacillus larvae, 442–443 Paenibacillus lentimorbus, 442 Paenibacillus macerans, 443, 453 Paenibacillus pasadenensis, 443 Paenibacillus polymyxa, 442–443, 449, 452– 453 Paenibacillus popilliae, 442–443 Paenibacillus provencensis, 443 Paenibacillus sanguinis, 443 Paenibacillus thiaminolyticus, 443 Paenibacillus timonensis, 443 Paenibacillus turicensis, 443 Paenibacillus urinalis, 443 Paenibacillus validus, 453 Paenibacillus vulneris, 443 Page’s ameba saline, 2393 Pain, see also specific locations arenaviruses, 1673–1674 filoviruses, 1674 Pairwise analysis of sequence conservation (PASC), 1395, 1397 Pajaroello, 2515 Pajuello, 2512 Palivizumab, 1501, 1508 Palivizumab resistance, 1508 P-Aminosalicylic acid (PAS), for Mycobacterium infection, 1361 Pampa virus, 1669, 1672 PANArray, 1792

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 120

Pancreatic abscess, Aeromonas, 754 Pancreatitis Ascaris lumbricoides, 2451 Campylobacter, 1000 Dolosigranulum pigrum, 424 liver trematodes, 2489 Pandemics, influenza, 1432, 1470–1471 Pandoraea, 617, 632–633 characteristics of, 798 clinical significance, 795 description of genus, 792 direct examination, 795 epidemiology and transmission, 793 identification, 797–799, 801 taxonomy, 791–792 typing systems, 803 Pandoraea apista, 632–633, 791 Pandoraea faecigallinarum, 792 Pandoraea norimbergensis, 632–633, 792 Pandoraea pnomenusa, 632–633, 791 Pandoraea pulmonicola, 632–633, 791 Pandoraea sputorum, 632–633, 791 Panniculitis, Gnathostoma and, 2497 Pannonibacter phragmitetus, 626–627, 823–824 Panstrongylus, 2507 Panther system, 74 Pantoea epidemiology, transmission, and clinical significance, 719 identification, 725 taxonomy, 714 Pantoea agglomerans, 715, 717, 719, 723, 725 Pantoea ananatis, 717, 719, 725 Pantoea brenneri, 717, 719 Pantoea citrea, 719 Pantoea conspicua, 717, 719 Pantoea cypripedii, 719 Pantoea dispersa, 719, 725 Pantoea eucrina, 717, 719 Pantoea punctata, 719 Pantoea septica, 717, 719 Pantoea stewartii, 719 Pantoea terrea, 719 Panton-Valentine leukocidin (PVL), 366 Pap smears, human papillomavirus and, 1785 Papanicolaou stain, for fungi, 1970 Paper wasps, 2518 PapilloCheck, 1792, 1795 Papilloma mother yaw, 1061 viruses, specimens and methods for detection of, 1406 Papillomaviridae (family), 1398, 1400–1401 Papillomaviruses, see also Human papillomavirus detection and identification methods, 1434 specimen collection and handling, 1406, 1408 Papovaviridae (family), 1803 PapType HPV test, 1792 Papules, viral, 1406, 1408 Papulosa stomatitis virus, 1828, 1830 para-Aminobenzoic acid, 1191 Parabacteroides antimicrobial susceptibilities, 984, 1185, 1346 characteristics of genus, 970–971 clinical significance, 971 identification, 977–978 taxonomy, 967, 969 Parabacteroides distasonis, 967, 970–971, 978, 983–984, 1346, 1350 Parabacteroides goldsteinii, 967, 971, 977–978

Parabacteroides gordonii, 967, 971, 977–978 Parabacteroides johnsonii, 967, 978 Parabacteroides merdae, 967, 971, 977–978 Parabuthus, 2520 Paracoccidioides, 2196 Paracoccidioides brasiliensis, 1935, 1939, 2094, 2109–2123 antifungal susceptibilities, 2121–2122, 2224 antigen detection, 2116 biosafety, 2117 clinical significance, 2114–2115 culture for mold phase, 2117 culture for yeast phase, 2118 description of agents, 2112 direct examination, 2115–2117 endophthalmitis, 1949 epidemiology and transmission, 2113–2114 evaluation, interpretation, and reporting of results, 2123 identification, 2119 isolation, 2117–2118 Lacazia loboi, compared to, 2196–2199 media, 1959 microscopy, 1966, 1976, 2112, 2116 nucleic acid detection, 2117 serologic tests, 2121 specimen collection, transport, and processing, 1948–1949, 1951, 2115 taxonomy, 2110 typing systems, 2120 Paracoccidioides lutzii, 2110 Paracoccidioidomycosis, 2109–2123 antigen detection, 2116 clinical significance, 2114–2115 description of agent, 2112 epidemiology and transmission, 2113–2114 evaluation, interpretation, and reporting of results, 2123 nucleic acid detection, 2117 serologic tests, 2121 specimens for, 1947, 1950, 2115 Paracoccus yeei, 632–633, 820, 822 Paraeggerthella clinical significance, 925 taxonomy and description, 920–921 Paraeggerthella hongkongensis, 925, 930 Paraffin oil, storage of microorganisms in, 166 Paraformaldehyde, 193 Parafungi, 1936 Paragonimiasis, 2479, 2507, 2533 Paragonimidae (family), 2290 Paragonimus, 2479 arthropod vector, 2507, 2513 detection, 2320, 2329, 2331 sputum specimen, 2305 treatment, 2531 Paragonimus africanus, 2481 Paragonimus caliensis, 2481, 2487 Paragonimus heterotremus, 2481, 2487 Paragonimus hueitungensis, 2481, 2487 Paragonimus kellicotti, 2481, 2487 Paragonimus mexicanus, 2481, 2487 Paragonimus miyazakii, 2481, 2487 Paragonimus uterobilateralis, 2481, 2487 Paragonimus westermani, 2449, 2481, 2484, 2487–2488 Parainfluenza virus, 1487–1492 antigen detection, 1489 clinical significance, 1488 cytopathic effect (CPE), 1491 description of agents, 1487 detection and identification methods, 1434

SUBJECT INDEX DFA and IFA reagents for the detection of, 1425 diagnostic methods for detection, 1489 direct examination, 1488–1491 epidemiology and transmission, 1487 evaluation, interpretation, and reporting of results, 1492 immunofluorescence detection in R-Mix cells, 1426 in immunocompromised patients, 1488 isolation and identification, 1491 microscopy, 1488 nucleic acid detection, 1489–1491 rapid cell culture, 1426 serologic tests, 1492 specimen collection and handling, 1407– 1408, 1488 taxonomy, 1487 vaccines, 1488 Paralysis enterovirus, 1540 herpes B virus, 1697 herpes simplex virus (HSV), 1689 parechovirus, 1541 poliomyelitis, 1538–1540 rabies virus, 1635 spider envenomation, 2520 tick paralysis, 2516 Paramphistomidae (family), 2482 Paramphistomoidea (superfamily), 2290 Paramyxoviridae (family), 1398, 1400, 1401, 1487, 1498, 1508, 1519 Paramyxovirinae (subfamily), 1398, 1487 Parana virus, 1669, 1671 Parapoxvirus (genus), 1398, 1828–1829, 1834–1835 Parapoxviruses, 1828–1837, 1835 antigen detection, 1833 clinical significance, 1831 collection, transport, and storage of specimens, 1832 description of agents, 1828 diagnostic tests, 1832 direct detection, 1832–1835 epidemiology and transmission, 1828–1830 evaluation, interpretation, and reporting of results, 1837 identification, 1835 isolation, 1835 microscopy, 1832–1833 nucleic acid detection, 1833–1835 serologic tests, 1837 taxonomy, 1828–1829 Paraprevotella, 968 Paraprevotella clara, 968 Paraprevotella xylaniphila, 968 Parasafe, 2311 Parascardovia, 920–921 Parascardovia denticolens, 925 Parasite, definition of term, 2285 Parasite lactate dehydrogenase (pLDH) test, 2336 Parasitiformes (order), 2511 Parasitology antiparasitic agents, 2529–2545 resistance mechanisms, 2550–2556 susceptibility testing methods, 2563– 2568 blood sample, 2297, 2304, 2306–2307, 2333–2336 antigen and DNA detection, 2307 blood stains, 2306 buffy coat films, 2307, 2336 collection, 2304

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 121

concentration procedures, 2307, 2336 detection and identification, 2333–2336 examination of films, 2335 immunochromatographic tests for malaria, 2335–2336 Knott concentration, 2307, 2336 membrane filtration, 2307, 2336 preparation of films, 2333–2334 screening methods, 2307 staining, 2334–2335 thick blood films, 2306, 2333–2335 thin blood films, 2306, 2333–2335 detection and identification, 2317–2337 blood, 2333–2336 bone marrow, 2327–2330 eyes, 2328, 2330 intestinal tract specimens, 2324–2326 kidneys and bladder, 2328, 2330 liver and spleen, 2328, 2330–2331 lungs, 2329, 2331 lymph nodes and lymphatics, 2328, 2331–2332 muscle, 2329, 2332 nasopharynx and sinus cavities, 2329, 2332 rectal tissue, 2329, 2332 skin, 2329, 2332–2333 stool specimens, 2317–2324 urogenital specimens, 2326–2327 direct detection by routine methods, 2304–2308 amniotic fluid, 2305 animal inoculation, 2307 antigen detection, 2307–2308 aspirates, 2305–2306 biopsy specimens, 2306 blood, 2306–2307 culture methods, 2307 intestinal tract specimens, 2304–2305 sputum, 2305 urogenital tract specimens, 2305 xenodiagnosis, 2307 media, 2315–2316 reagents, 2310–2312 risk-based classification, 171 specimen collection, transport, and processing, 2293–2308 blood collection, 2304 body sites and possible parasites recovered, 2294 commercial kits for immunodetection in stool samples, 2295 commercial kits for immunodetection of serum antibodies, 2296 direct detection by routine methods, 2304–2308 fecal specimen collection and processing options, 2301 sample preparation and procedures by body site, 2297 stool collection, 2293–2294, 2296 stool preservation, 2300–2304 stool test ordering, 2302 stains, 2312–2314, 2316 stool specimens, 2293–2304, 2317–2324 concentration wet mount, 2317–2318 culture of larval-stage nematodes, 2321– 2323 direct wet mount in saline, 2317 egg identification, 2320 hatching of schistosome eggs, 2323 helminth recovery and identification techniques, 2323 immunoassay methods, 2319–2320, 2322

n cxxi

key to identification of intestinal amebae, 2321 key to identification of intestinal flagellates, 2322 molecular methods, 2320–2321 permanent stained smears, 2318–2319 processing liquid stool, 2318 processing preserved stool, 2318–2319 taenia solices, search for, 2324 worm burden estimation, 2323 taxonomy and classification, 2285–2291 acanthocephalans, 2291 cestodes, 2288, 2291 helminths, 2288–2291 nematodes, 2288–2289 protozoa, 2285–2288 trematodes, 2288, 2290 Parasitosis, delusion or illusion of, 2521 Parastrongylus, 2498–2499 clinical significance, 2498 description of agents, 2498 direct examination by microscopy, 2498 epidemiology, transmission, and prevention, 2498 serologic tests, 2498–2499 treatment, 2499 Parastrongylus cantonensis, 2498–2499 Parastrongylus costaricensis, 2498 Parasutterella, 969, 981 Parasutterella excrementihominis, 969, 981 Parasutterella secunda, 969, 981 PARA-TECT Cryptosporidium/Giardia DFA, 2295 PARA-TECT Cryptosporium, 2295 PARA-TECT Giardia, 2295 Paravaccinia, 1829 Paravahlkampfia, 2387, 2392 Paravahlkampfia francinae, 2387 parC Aeromonas, 1326 Neisseria gonorrhoeae, 1323, 1383 Vibrio cholerae, 1331 Parechovirus (genus), 1399, 1551 Parechoviruses, 1536–1546 antiviral susceptibilities, 1545 cell lines, susceptible, 1543–1544 clinical significance, 1538–1541 cytopathic effect, 1543–1544 detection and identification methods, 1433 direct examination, 1542–1543 epidemiology and transmission, 1537–1538 genome organization, 1537 identification, 1543–1544 isolation procedures, 1543 nucleic acid detection, 1542–1543 serologic tests, 1545 serotypes, 1537 specimen collection and handling, 1406– 1407 typing systems, 1545 Paresthesia, polymyxins and, 1193 Parinaud’s oculoglandular syndrome, 876 Paritaprevir, 1879–1880 Parker ink stain, 1973 Parkinson’s disease, 1018 Paromomycin, 2543–2544 Acanthamoeba, 2394 adverse effects, 2544 Cryptosporidium, 2442 Dientamoeba fragilis, 2413 Entamoeba histolytica, 2405 Giardia duodenalis, 2412 Leishmania, 2361–2362

cxxii

n

SUBJECT INDEX

Paromomycin (continued) leishmaniasis, 2564 mechanism of action, 2543 pharmacokinetics, 2543 spectrum of activity, 2543–2544 Trichomonas vaginalis, 2554 Parotitis mumps virus, 1493 Staphylococcus, 360 viruses, specimens and methods for detection of, 1406 Particle agglutination assay human T-cell lymphotropic viruses (HTLVs), 1462–1463 Mycoplasma, 1097 Parvimonas, 909, 1343 Parvimonas micra, 909–915 Parvoviridae (family), 1398, 1400–1401, 1618, 1818, 1823 Parvovirinae (subfamily), 1398, 1818 Parvovirus detection and identification methods, 1434 taxonomy, 1818 Parvovirus 4 (PARV4), 1818, 1824 Parvovirus B19, 1818–1823 antigen detection, 1821 clinical significance, 1818–1829 collection, transport, and storage of specimens, 1406–1407, 1412, 1820–1821 description, 1818 direct examination, 1821 epidemiology and transmission, 1818 evaluation, interpretation, and reporting of results, 1822–1823 isolation, 1822 microscopy, 1821 nucleic acid detection, 1819, 1821–1822 serologic tests, 1822 taxonomy, 1818 time course of infection, 1820 treatment, 1820 typing systems, 1822 Paryphostomum, 2482 Pasteurella, 652 antimicrobial susceptibilities, 661–662 antimicrobial susceptibility testing, 1318, 1331 clinical significance, 655 direct examination, 656 epidemiology and transmission, 654 identification, 658, 660–661 isolation procedures, 656 serotyping, 661 taxonomy and description of, 653 Pasteurella aerogenes, 654–655, 661 Pasteurella bettyae, 654–655, 661 Pasteurella caballi, 654–655, 661 Pasteurella canis, 653, 655, 661 Pasteurella dagmatis, 653, 655, 661 Pasteurella haemolytica selective medium, 342 Pasteurella multocida, 653, 655, 661–662 antimicrobial susceptibilities, 1172, 1183, 1197 differentiation of Francisella from, 852 Pasteurella multocida subsp. gallicida, 653 Pasteurella multocida subsp. multocida, 653, 655 Pasteurella multocida subsp. septica, 653, 655 Pasteurella pneumotropica, 654–655, 661 Pasteurella stomatis, 653, 655, 661 Pasteurellaceae (family), 652–653, 667 Pasteuria, 441–442

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 122

Pasteuriaceae, 441 Pastorex Candida, 1996 Pastorex Crypto Plus, 1997 Pastorex Rotavirus, 1624 Pastorex Staph Plus, 363 PATH (Prospective Antifungal Therapy) Alliance, 1992 PathChip, 241 PathoDx respiratory virus panel, 1473 PathoDx RSV and respiratory virus panel, 1503 Pathogen discovery, 238–247 history classical methods, 238 microbial genomics, 240–241 molecular strategies, 238, 240 timeline, 239 microbial genomics microarray-based approaches, 241–242 recent history of, 240–241 sequencing-based approaches, 242–246 Pathogen evolution, 147 PathogenFinder, 246 Pathogenicity, role of, 246–247 Pathogenicity island, Enterococcus, 405 Pathozyme Syphilis, 1068, 1071 Patulin, 2189–2191 Pauciseptate, 1937, 1942 PCMX (chloroxylenol), 186 PCR (polymerase chain reaction), see also specific PCR methods Acinetobacter, 819 adenovirus, 1774–1775, 1778 amebae, pathogenic and opportunistic free-living, 2392–2393 Anaplasma phagocyrophilum, 1143 Anisakis, 2493 antibiotic resistance detection in staphylococci oxacillin resistance, 1293–1294 penicillin resistance, 1289 antimicrobial susceptibility testing, 1247 arboviruses, 1648–1652 Arcobacter, 1005–1006 arenaviruses, 1676–1677, 1681 Aspergillus, 2039–2040 Babesia, 2351–2352 Bacillus anthracis, 1325 Bartonella, 877–880 Bordetella, 842 Borrelia, 1042–1043 Brachyspira, 1064 Brucella, 867 Burkholderia pseudomallei, 796 Campylobacter, 1005–1006 Candida, 1997 capillary electrophoresis integrated with, 67 Chlamydiaceae, 1110–1112 classification and identification of bacteria and, 261 Clostridium, 956 Clostridium perfringens, 948–949 coccidia, 2430 competitive (cPCR), 73 coronaviruses, 1567, 1569–1577 Coxiella burnetii, 1153–1155 Cryptosporidium, 2440–2441 cytomegalovirus, 1723–1726 dermatophytes, 2139, 2141 Dientamoeba fragilis, 2413 digital PCR, 63, 64 dimorphic fungi causing systemic mycoses, 2117

diphtheria toxin testing, 488 DNA microarrays and, 144 Ehrlichia chaffeensis, 1140 Entamoeba histolytica, 2405–2406 Enterobacteriaceae, 726 enteroviruses, 1542–1543 entomophthoromycosis, 2100 Epstein-Barr virus, 1742–1744 eumycotic mycetoma, 2178 filoviruses, 1676–1677, 1681 Francisella tularensis, 856–859 fungal identification and diagnosis, 1979 Fusarium, 2068 gastroenteritis viruses, 1625–1627 Giardia duodenalis, 2412 hantaviruses, 1663, 1665 Helicobacter, 1019–1021, 1023–1024 hepatitis C virus, 1603–1607, 1611 herpes B virus, 1697 herpes simplex virus (HSV), 1690–1691, 1695–1696 Histoplasma capsulatum, 2118 human bocavirus, 1824 human herpesvirus 6 (HHV-6), 1756– 1757, 1759–1760 human herpesvirus 7 (HHV-7), 1758– 1761 human herpesvirus 8 (HHV-8), 1758– 1760, 1763–1764 human immunodeficiency virus, 1441– 1443, 1447, 1450–1451 human T-cell lymphotropic viruses (HTLVs), 1461–1462, 1462 immuno-PCR, 102–103 influenza viruses, 1476–1478, 1480–1482 Legionella pneumophila, 892, 898 Leishmania, 2359, 2361 Leptospira, 1031–1032 Loa loa, 2467 lymphatic filarial nematodes, 2465 MALDI-TOF MS combined with, 72–73 Mansonella, 2468 measles virus, 1522, 1525 melanized fungi, 2164 molecular detection of antibacterial resistance, 1379–1380, 1383, 1385 mucormycosis, 2090–2091 Mycobacterium tuberculosis complex, 1367– 1368 Mycoplasma, 1088–1098, 1100–1101 Onchocerca volvulus, 2466 overview, 57, 59 parainfluenza virus, 1490 parasites, 2297–2300, 2308 parvovirus B19, 1819, 1821–1823 pathogen discovery and, 238, 240 Plasmodium, 2347–2348 Pneumocystis, 2024–2025 polyomaviruses, 1809–1810 poxviruses, 1833–1835 Pseudomonas, 780 rabies virus, 1637, 1640 respiratory syncytial virus, 1502, 1506– 1508, 1510–1512 reverse transcriptase PCR (RT-PCR), 57– 58 Rickettsia, 1124, 1126–1128, 1130 rubella virus, 1527–1528, 1530 schistosomes, 2486 Stenotrophomonas maltophilia, 801–802 subtyping methods random amplification of polymorphic DNA/arbitrarily primed PCR (RAPD/AP-PCR), 137

SUBJECT INDEX repetitive element PCR (rep-PCR), 137 RFLP combined with, 138–139 ribotyping, 137 Taenia saginata, 2474 Taenia solium, 2475 Talaromyces marneffei, 2046 Toxoplasma gondii, 2375–2376 Treponema, 1062–1065, 1073 Trichomonas vaginalis, 2415 Tropheryma whipplei, 1160–1165 Trypanosoma brucei, 2367 Trypanosoma cruzi, 2365 varicella-zoster virus, 1706–1710 PCR electrospray ionization mass spectrometry, 39–40, 73, 1380, 1571 PCR ribotyping, 137, 956 PCR-restriction enzyme analysis, Mycobacterium, 603 p-Dimethylaminocinnamaldehyde (DMACA), 318 Pectobacterium cypripedii, see Pantoea cypripedii Pediatric blood culture bottles, 22 Pediatric infections, Enterobacteriaceae and, 715 Pediculicide, 2511 Pediculosis, 2510–2511 Pediculus, 2507 Pediculus humanus capitis, 2330, 2510–2511 Pediculus humanus corporis, 1126, 2330, 2510–2511 Pediculus humanus humanus, 1037–1038, 1040 Pediococcus antimicrobial susceptibilities, 430, 1184, 1189–1190 antimicrobial susceptibility testing, 1318, 1330 clinical significance, 424 epidemiology and transmission, 423 identification, 426–427, 429 interpretation of results, 431 isolation procedures, 425 taxonomy, 422 Pediococcus acidilactici, 422, 424, 427, 430 Pediococcus halophilus, 422 Pediococcus pentosaceus, 422, 424, 427 Pefloxacin, 1178, 1199 Pegylated interferon alpha hepatitis B virus, 1881–1882, 1900 hepatitis C virus, 1878–1879, 1901 Pegylated interferon α2a resistance, 1851 PEL (primary effusion lymphoma), 1763– 1764 Pelczaria aurantia, see Kocuria rosea Pelvic abscess, 290 Pelvic inflammatory disease (PID) Chlamydia trachomatis, 1108 Mycoplasma, 1092 non-spore-forming, anaerobic, Grampositive rods, 923 Ureaplasma, 1092 Pemphigus neonatorum, 360 penA gene Neisseria gonorrhoeae, 1322, 1383 Neisseria meningitidis, 1323–1324 Penciclovir herpes simplex virus (HSV), 1689, 1919 herpesviruses, 1884 varicella-zoster virus, 1706, 1712 Penciclovir resistance, 1917 herpes simplex virus (HSV), 1894–1895 varicella-zoster virus, 1712, 1895 Penicillin(s), 1171–1173 adverse effects, 1173

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 123

anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1255, 1259 antistaphylococcal, 1171 carboxypenicillins, 1173 concentration in serum, 1198–1199 extended spectrum, 1172 isoxazolyl, 1171 mechanism of action, 1171 penicillinase-resistant, 1172 pharmacology, 1171 repository forms, 1171 semisynthetic, 1172 spectrum of activity, 1171–1173 structure, 1172 ureidopenicillins, 1173 Penicillin G, 1171–1173, 1199, 1255, 1259 Penicillin resistance, 1172–1173 Bacillus, 1326 Bacteroides fragilis group, 1346 detection by automated antimicrobial susceptibility testing, 1278 Fusobacterium, 1348 in staphylococci, 1278, 1289 Neisseria gonorrhoeae, 1322 Neisseria meningitidis, 1323–1324 Prevotella, 1347 Streptococcus, 1320 Streptococcus pneumoniae, 1315–1316 Penicillin V, 1171–1172, 1199 Penicillin zone edge test, 1302 Penicillinase-producing Neisseria gonorrhoeae, 324 Penicillinase-producing Neisseria gonorrhoeae medium (PPNG selective medium), 342 Penicillinase-resistant penicillins, 1172 Penicillinases, 1172, 1177, 1224–1226, 1289 Penicillin-binding proteins (PBPs) carbapenems and, 1176 cephalosporins and, 1173 Haemophilus influenzae, 1321 monobactams and, 1176 Neisseria gonorrhoeae, 1322 Neisseria meningitidis, 1323 PBP-mediated β-lactam resistance, 1212, 1220–1223, 1291 acquisition of foreign PBPs, 1221–1222 PBP overexpression, 1221 point mutations, 1222–1223 resistance mutations by recombination with foreign DNA, 1222 penicillins and, 1171 Streptococcus pneumoniae, 1315, 1383 Penicilliosis, specimens for, 1947 Penicillium, 1171, 1938, 1939, 1940, 2045– 2048, 2230 citrinin, 2189 clinical significance, 2048 cyclopiazonic acid, 2189 identification, 2048 sick building syndrome, 2192 taxonomy, 2045–2046, 2048 Penicillium aurantiogriseum, 2048, 2189 Penicillium brevicompactum, 2048 Penicillium camemberti, 2189 Penicillium chrysogenum, 2048 Penicillium citrinum, 2047–2048, 2189 Penicillium commune, 2048 Penicillium decumbens, 2048 Penicillium expansum, 2048, 2189 Penicillium griseofulvum, 2048, 2190, 2230 Penicillium janthellum, 2047

n cxxiii

Penicillium marneffei, 2115 Penicillium purpurogenum, 2047 Penicillium rubrum, 2047 Penicillium spinulosum, 2048 Penicillium verrucosum, 2189 Penile human papillomavirus, 1784–1785 Pentamidine, 2543 adverse effects, 2543 African trypanosomiasis, 2564 Leishmania, 2361 leishmaniasis, 2564 mechanism of action, 2543 pharmacokinetics, 2543 spectrum of activity, 2543 Pentamidine isethionate Pneumocystis, 2025–2026 Trypanosoma brucei, 2367 Pentamidine isothiocyanate, for Balamuthia mandrillaris, 2395 Pentamidine resistance, 2551, 2555 Pentastomes, 2516 Pentatrichomonas hominis, 2321, 2400, 2408, 2410, 2414, 2416 Pentavalent antimonials, 2542 adverse effects, 2542 Leishmania, 2361–2362 mechanism of action, 2542 pharmacokinetics, 2542 spectrum of activity, 2542 Pentavalent antimonials resistance, 2551, 2554 Pentostam, 2542 Peptic ulcer disease, Helicobacter and, 1017 Peptide nucleic acid (PNA) probes, 55 Peptide nucleic acid-fluorescent in situ hybridization (PNA-FISH), 23, 1381 Pseudomonas, 776 yeasts, 1997 Peptococcus, 909 Peptococcus niger, 909–910, 1348 Peptococcus saccharolyticus, see Staphylococcus saccharolyticus Peptone iron agar, 342 Peptone-yeast extract-glucose (PYG) medium, 2315 Peptoniphilus, 909, 911, 1348 Peptoniphilus asaccharolyticus, 909–915 Peptoniphilus coxii, 909–910, 913–914 Peptoniphilus duerdenii, 909–910, 914 Peptoniphilus gorbachii, 909–910, 913–915 Peptoniphilus grossensis, 909 Peptoniphilus harei, 909–911, 914–915 Peptoniphilus indolicus, 909–911, 914 Peptoniphilus ivorii, 909–911, 914–915 Peptoniphilus koenoeneniae, 909–910, 914 Peptoniphilus lacrimalis, 909–910, 914–915 Peptoniphilus methioninivorax, 909 Peptoniphilus olsenii, 909–910, 914 Peptoniphilus timonensis, 909 Peptoniphilus tyrrelliae, 909–910, 913–914 Peptostreptococcus, 909–916 antimicrobial susceptibilities, 913, 916, 1175, 1183–1184, 1189–1190 clinical significance, 911 description of, 909 direct examination, 912 epidemiology, 909–910 identification, 912–915 taxonomy, 909 Peptostreptococcus anaerobius, 909–915 Peptostreptococcus magnus, 909 Peptostreptococcus micros, 909, 913 Peptostreptococcus morbillorum, 422 Peptostreptococcus productus, 909, 914–915; see also Blautia producta

cxxiv

n

SUBJECT INDEX

Peptostreptococcus russellii, 914 Peptostreptococcus saccharolyticus, see Staphylococcus saccharolyticus Peptostreptococcus stomatis, 909–911, 913–916 Peracetic acid, 194 Peramivir, 1886–1887 Percolozoa (phylum), 2287 Perenniporia, 2071, 2075 Perfringens agar (Shahidi-Ferguson perfringens agar), 342 Pergolide, 2190 Pericardial effusion, arenavirus, 1673 Pericardial fluid specimens collection, transport, and storage guidelines, 276 fungi, 1946, 1950 Pericarditis anaerobic Gram-negative rods, 972 enterovirus, 1541 Fusobacterium, 973 Gram-positive anaerobic cocci (GPAC), 910 Mansonella, 2468 measles, 1521 Mycobacterium kansasii, 542 Mycobacterium tuberculosis, 538 Mycoplasma, 1091 viruses, specimens and methods for detection of, 1406 Perihepatitis, Chlamydia trachomatis, 1108 Peri-implant disease, anaerobic Gramnegative rods and, 972 Periodic acid-Schiff (PAS) stain, 1958 fungi, 1970 Tropheryma whipplei, 1159–1160, 1162 Periodontal disease/infection, 229 Aggregatibacter actinomycetemcomitans, 654 anaerobic Gram-negative rods, 969, 972, 974 Atopobium, 925 Campylobacter, 1001 Capnocytophaga, 653 Cryptobacterium curtum, 925 Desulfovibrio, 974 Dialister, 974 Eubacterium, 924 Fusobacterium, 973 Kingella, 655 non-spore-forming, anaerobic, Grampositive rods, 923 Olsenella, 925 Peptostreptococcus anaerobius, 911 Porphyromonas, 971 Prevotella, 972 Selenomonas, 974 Slackia exigua, 925 Tannerella forsythia, 971 Treponema, 1058–1059, 1063 Periorbital cellulitis, 290 Perirectal abscess, Sutterella, 974 Perithecium, 1942, 2070 Peritoneal fluid specimens collection, transport, and storage guidelines, 276 fungi, 1946, 1950 Peritonitis Abiotrophia and Granulicatella, 424 Achromobacter, 841 Aerococcus, 424 Aeromonas, 754 anaerobic Gram-negative rods, 972 Arcobacter, 1001 Asaia, 829 Bacillus cereus, 443

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 124

Bacillus circulans, 443 Bacteroides, 970 Campylobacter, 1000–1001 Capnocytophaga, 654 Clostridium, 948 Entamoeba histolytica, 2403 Fusarium, 2058 Gemella, 424 Haemophilus haemolyticus, 670 Lactococcus, 424 liver trematodes, 2489 Methylobacterium, 830 microsporidia, 2210, 2213 Neisseria gonorrhoeae, 636 non-spore-forming, anaerobic, Grampositive rods, 923 Pasteurella, 655 phaeohyphomycoses, 2163 Porphyromonas, 971 Pseudomonas, 776 Rhodotorula, 1994 Rothia mucilaginosa, 361 Shewanella, 825 Sphingobacterium, 825 Stenotrophomonas maltophilia, 794 Sutterella, 974 Tsukamurella paurometabola, 519 vancomycin-resistant lactobacilli, 924 Vibrio fluvialis, 765 Vibrio metschnikovii, 766 Perkinsus, 2287 Permanent stained specimens amebae, pathogenic and opportunistic free-living, 2392 sigmoidoscopy material, 2324–2325 sputum specimen, 2331 stool specimen, 2318–2319 Permethrin, 2516–2517 Permount, 2334 Pernicious anemia, 2472–2473 Peroxygen compounds, 195 Persister cells, azole resistance and, 2243 Personal protective equipment, 175, 283 Pertussis, 125, 841, 845 Petechiae arenaviruses, 1674 filoviruses, 1674 Petragnani medium, 342 Petriella, 2159 Petrolatum, 2518 Petromyces, 2030 Petromyces alliaceus, 2031–2032 Petromyces flavus, 2032, 2034 Petromyces nominus, 2031 PFGE, see Pulsed-field gel electrophoresis Pfizer TB medium base with glycerol, egg yolk, glucose, and malachite green, 342 PGT medium, 342 pH indicators, 320, 321 Phadebact Monoclonal GC test, 641 Phaeoacremonium, 2153–2155, 2158–2159, 2162, 2165, 2173–2174, 2177–2179, 2181 Phaeoacremonium alvesii, 2154 Phaeoacremonium griseobrunneum, 2155 Phaeoacremonium inflatipes, 2173, 2177 Phaeoacremonium krajdenii, 2155, 2173, 2178 Phaeoacremonium parasiticum, 1936, 2071, 2155, 2157, 2159, 2162, 2173 Phaeoacremonium rubrigenum, 2155 Phaeoacremonium venezuelense, 2155 Phaeoannellomyces werneckii, 2147 Phaeohyphomycosis, 1969, 2057, 2153

antifungal susceptibilities, 2167 clinical significance, 2161–2163 epidemiology and transmission, 2161 microscopy, 2164 Phage display, 246 Phagicola, 2482 Phanerochaete chrysosporium, 2063, 2071 Pharyngitis adenoviruses, 1771, 1772 Arcanobacterium haemolyticum, 479 Chlamydia pneumoniae, 1108 Corynebacterium diphtheriae, 479, 1327 cytomegalovirus, 1719 Epstein-Barr virus, 1739 herpes simplex virus (HSV), 1688 Mycoplasma, 1091 Neisseria gonorrhoeae, 636 S. pyogenes, 397 specimen collection, transport, and handling, 299–300 Streptococcus pyogenes, 385, 389 viruses, specimens and methods for detection of, 1408 Pharyngoconjunctival fever, adenovirus, 1772 Pharynx specimen collection, transport, and storage guidelines, 279 Phascolarctobacterium succinatutens, 969 Phase-contrast microscopy, 9 Phase Lock system, 2024 Phasmidea (class), 2289 Phenol, 194 Phenol oxidase test, for yeasts, 2000 Phenol red, 319 Phenol red agar, 342 Phenol red tartrate broth, 342 Phenolics, 194–195 PhenoSense assay, 1448 Phenotypic characteristics of bacteria, for classification and identification, 261– 262 Phenotypic identification systems, 32–35 Phenotypic methods for detecting antibacterial resistance, 1286–1303 direct tests for β-lactamases, 1302–1303 in Enterobacteriaceae, 1287, 1298–1302 in enterococci, 1286–1289 in staphylococci, 1287, 1289–1297 in streptococci, 1297–1298 quality control, 1286, 1288–1289, 1291, 1295–1298, 1303 Phenylalanine deaminase test, 319 Phenylethyl alcohol agar (phenylethanol agar, phenylethyl alcohol agar), 342 Phialemoniopsis, 2064, 2069, 2071, 2076 Phialemoniopsis cornearis, 2064, 2071 Phialemoniopsis curvata, 2064, 2071, 2075, 2076, 2157 Phialemoniopsis curvatum, 2161 Phialemoniopsis dimorphosporum, 2071 Phialemoniopsis ocularis, 2064, 2071 Phialemoniopsis pluriloculosa, 2064, 2071 Phialemonium, 2064, 2069, 2070, 2071, 2076, 2153, 2161 Phialemonium atrogriseum, 2064, 2071 Phialemonium curvatum, 2071, 2076, 2161 Phialemonium globosum, 2064, 2071 Phialemonium inflatum, 2064, 2071 Phialemonium obovatum, 2064, 2071, 2076, 2155 Phialides, 1940, 1942, 2058, 2070 Phialidic conidiogenesis, 1940 Phialophora, 1939, 1940, 2153–2154, 2158, 2271

SUBJECT INDEX Phialophora americana, 2154, 2158, 2163 Phialophora europaea, 2154, 2161, 2162 Phialophora parasitica, 1936 Phialophora richardsiae, 2155 Phialophora verrucosa, 1967, 2154, 2158, 2163 Phialosimplex, 2064, 2073, 2076 Phialosimplex caninus, 2064, 2073, 2075, 2076 Phialosimplex chlamydosporus, 2064, 2073 Phialosimplex sclerotialis, 2064, 2073 Phlebitis, streptogramins and, 1190 Phlebotomus, 2507 Phlebovirus, 1651, 2507 Phlebovirus (genus), 1399, 1645 Phocaeicola, 981–982 Phocaeicola abscessus, 981–982 Phoenix instrument, 33–34, 428, 1277; see also BD Phoenix system Phoma, 2065, 2069, 2075, 2076 Phomopsis, 2190 Phoneutria, 2520 Phoneutrism, 2520 Phormia regina, 2517 Phosphate-buffered saline, 320 Phosphotransferases, 1220, 1383 Photobacterium, 762, 763 Photobacterium damselae, 762, 764–765, 768– 769, 996–997 Photobleaching (fading), 11 Photomicroscopy, 11–12 Photophobia arboviruses, 1647 arenaviruses, 1674 herpes simplex virus (HSV), 1689 Pythium insidiosum, 2201 Photorhabdus asymbiotica, 715, 717, 722, 725– 726 Photorhabdus asymbiotica subsp. asymbiotica, 717 Photorhabdus asymbiotica subsp. australis, 717 Photorhabdus luminescens, 715, 717 Photorhabdus temperata, 717 Photosensitivity quinolones, 1180 tetracyclines, 1187 Phthiraptera, 2510–2511 Phthirus pubis, 2510–2511 PhyloChip, 242 Phylogenetic approach to species recognition (PSR concept), 1937 Phylogeny, 131 Physalopteridae (family), 2289 Physalopteroidea (superfamily), 2289 Phytoplasma, 1089 Pichia, 1938, 1985, 1988, 1993, 1998, 2001, 2263 Pichia angusta, 1993 Pichia anomala, 1990, 1993, 2005 Pichia fabianii, 1993 Pichia farinosa, 1993 Pichia kudriavzevii, 1985, 2000 Pichia norvegensis, 2000 Pichinde virus, 1669, 1671 Picobirnaviridae (family), 1399–1401, 1618 Picobirnavirus (genus), 1399, 1618 Picornavirales (order), 1395–1396, 1399, 1402 Picornaviridae (family), 1396, 1399–1401, 1536, 1545, 1551, 1584 Picornavirus (genus), 1558 PID, see Pelvic inflammatory disease Piedraia hortae, 2148 Pike streptococcal broth, 342 Pilonidal cyst, 278 Pinkeye, see Conjunctivitis

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 125

Pink-pigmented GNF, identification of, 827, 829–830 Pinta, 1055–1056, 1058, 1061 Pinworm, 2324; see also Enterobius vermicularis Piperacillin, 1173 antimicrobial susceptibility testing, 1255, 1259 concentration in serum, 1199 with tazobactam, 1178, 1199 Piperacillin-tazobactam anaerobic bacterial susceptibility percentages, 1351 antimicrobial susceptibility testing, 1255, 1259 Bacteroides fragilis group susceptibility percentages, 1350 Piperacillin-tazobactam resistance, 1346 Piperaquine, 2536, 2564 Piperaquine resistance, 2552 Piperazine, for Enterobius vermicularis, 2454 Pirital virus, 1669, 1672 Piroplasmida (order), 2287, 2338 Piry virus, 1645 Pityriasis folliculorum, 2517 Pityriasis versicolor, 1949, 1994, 2146–2147 Pityrosporum furfur, 2146 Pityrosporum orbiculare, 2146 Pityrosporum ovale pro parte, 2146 Placenta, sample handling, 292 Plagiorchiida (order), 2290 Plagiorchiidae (family), 2290 Plagiorchioidea (superfamily), 2290 Plague, 738–742; see also Yersinia pestis arthropod vector, 2507, 2509–2510 biothreat agent, 221 transmission and disease, 221 Planococcaceae, 441 Planococcus, 354–355 Plaque reduction assay (PRA) antiviral susceptibility testing, 1914–1916 Plaque reduction neutralization (PRN) measles virus, 1524 rubella virus, 1529–1530 Plaque reduction neutralization test (PRNT), for arboviruses, 1654–1655 Plasma sterilization, 204 Plasmid profiling, 135–136 Plasmids antibiotic resistance, 1216, 1230–1231, 1233–1235, 1250 Borrelia burgdorferi, 1040 Plasmodium antigen detection, 2307, 2347–2348 antimicrobial susceptibilities, 2349, 2536– 2541, 2545 antiparasitic agent resistance, 2550–2553 arthropod vector, 2507 blood specimens, 2306–2307 bone marrow aspirate, 2306 characteristics of infections, 2339 clinical significance, 2339–2341 collection, transport, and storage of specimens, 2341 description of agents, 2338 detection, 2327–2328, 2333–2336 direct examination, 2341–2348 epidemiology, 2338–2339 evaluation, interpretation, and reporting of results, 2349 Giemsa-stained blood films, 2341–2347 life cycle, 2340 media for culture, 2315–2316 microscopy, 2341–2348

n cxxv

morphology in Giemsa-stained thin films, 2343 nucleic acid detection, 2347–2348 serologic tests, 2349 storage methods, 166 taxonomy, 2338 Plasmodium falciparum, 2338–2349 antigen detection, 2307 antimicrobial susceptibilities, 1180, 1187, 2349, 2536–2539 antiparasitic agent resistance, 2550–2553 antiparasitic agent susceptibility testing methods, 2563–2566 Babesia morphology compared, 2352 characteristics of infections, 2339 clinical manifestations, 2341 detection, 2336 developmental stages, 2344 genome sequence, 241 morphology in Giemsa-stained thin films, 2343 Plasmodium knowlesi, 2338, 2341 characteristics of infections, 2339 developmental stages, 2346 morphology in Giemsa-stained thin films, 2343 treatment, 2536–2537, 2540–2541 Plasmodium malariae, 2338–2349 characteristics of infections, 2339 clinical manifestations, 2341 developmental stages, 2344 morphology in Giemsa-stained thin films, 2343 treatment, 2536–2537, 2539–2541 Plasmodium ovale, 2338–2349 antiparasitic agent resistance, 2553 characteristics of infections, 2339 clinical manifestations, 2341 developmental stages, 2345 morphology in Giemsa-stained thin films, 2343 treatment, 2536––2541 Plasmodium ovale curtisi, 2348 Plasmodium ovale wallikeri, 2348 Plasmodium vivax, 2338–2349 antiparasitic agent susceptibility testing, 2564–2565 characteristics of infections, 2339 clinical manifestations, 2341 developmental stages, 2345 morphology in Giemsa-stained thin films, 2343 treatment, 2536–2537, 2539–2541 Plate variability, EIA, 99 Platelia Aspergillus assay, 1971, 1997, 2038, 2044, 2067, 2116 Platelia Candida Ab test, 1996 Platelia Candida Ag test, 1972, 1996, 2067 Platyhelminthes (phylum), 2288, 2290, 2471, 2473–2475 Pleconaril, 1545, 1558 Pleistophora, 2209–2211, 2213, 2329, 2332 Pleistophora ronneafiei, 2209, 2213 Pleoanamorphism, 1942, 2153 Pleomorphic, 1942 Pleosporales (order), 2153, 2155, 2159–2163, 2173–2174, 2176 Plesiomonas antimicrobial susceptibilities, 1180 properties of, 763 Plesiomonas shigelloides, 715, 717, 764, 767 antimicrobial susceptibilities, 730 antimicrobial susceptibility testing, 1317, 1326

cxxvi

n

SUBJECT INDEX

Plesiomonas shigelloides (continued) collection, transport, and storage of specimens, 722 epidemiology, transmission, and clinical significance, 721 isolation procedures, 722 Pleural effusion, arenavirus, 1673 Pleural fluid specimens, 276 Pleuritis, Mycobacterium tuberculosis, 538 Pleurodesis, 1763 Pleurodynia enterovirus, 1540 viruses, specimens and methods for detection of, 1408 Pleurostomophora, 2153–2155, 2158 Pleurostomophora ochracea, 2154–2155, 2162, 2173–2174 Pleurostomophora repens, 2154–2155 Pleurostomophora richardsiae, 2154–2155, 2157, 2162 Pleurothecium obovoideum, 2159 Pleurotus mutilus, 1197 PLEX-ID, 145, 584, 1380–1381, 1383–1384 PLEX-ID broad fungal assay, 40 PLEX-ID flu, 1477 PLEX-ID respiratory viral panel, 1506, 1511 Plexus HerpeSelect 1 and 2 IgG kit, 1693– 1694 Pluralibacter, 715, 723 Pluralibacter gergoviae, 717, 719, 723 Pluralibacter pyrinus, 717, 719, 723 PML, see Progressive multifocal leukoencephalopathy PNA (peptide nucleic acid), Staphylococcus, 365 pncA gene, 1358–1359 Pneumococci, subtyping of, 139 Pneumocystidales (order), 1937 Pneumocystidomycetes (class), 1937–1938 Pneumocystis, 1936–1938, 2015–2026 antimicrobial susceptibilities, 185, 2025– 2026, 2543 clinical significance, 2018–2019 extrapulmonary pneumocystosis, 2019 high-resolution computerized tomography (HRCT), 2019 immune reconstitution inflammatory syndrome (IRIS), 2018 presentation in adults, 2019 presentation in children, 2018–2019 collection, transport, and storage of specimens, 2019–2022 bronchoalveolar lavage, 2020 induced sputum collection, 2020 nasopharyngeal aspirates, 2022 open thorax lung biopsy, 2020 oropharyngeal washes, 2022 transbronchial biopsy, 2022 colonization, 2018 description of agents, 2015–2016 developmental forms, 2016 direct examination and identification, 2022–2025 microscopy, 2022–2023 nucleic acid detection, 2024–2025 serum (1→3)-β-D-glucan, 2024 epidemiology and transmission, 2016–2018 evaluation, interpretation, and reporting of results, 2026 isolation, 2025 life cycle, 2016–2017 serologic tests, 2025 stains for detection, 2021–2022 taxonomy, 2015

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 126

Pneumocystis carinii, 2015–2016, 2026 Pneumocystis jirovecii, 1185, 1957–1958, 2015–2026, 2543 (1,3)-β-D-glucan detection, 1978 microscopy, 1966, 1975 reagents for, 1956 stains, 1970, 2316 Pneumocystis jirovecii pneumonia, 1440 Pneumocystis murina, 2015–2016 Pneumocystis oryctolagi, 2015 Pneumocystis pneumonia (PCP), 2016–2026 Pneumocystis wakefieldiae, 2015 Pneumocystosis, specimens for, 1947 Pneumonia Achromobacter, 841 Acinetobacter, 813 adenoviruses, 1771, 1772, 1773 Aeromonas, 754 Bacillus subtilis, 442 Balneatrix alpica, 827 BK polyomavirus, 1805 Blastomyces dermatitidis, 2114 Burkholderia, 794 Campylobacter, 1001 Candida, 2006 Chlamydia pneumoniae, 1109 Chlamydia psittaci, 1109 Chrysosporium zonatum, 2076 Coccidioides, 2114 coronaviruses, 1569 Dialister, 974 Dolosigranulum pigrum, 424 Elizabethkingia meningoseptica, 828 enterovirus, 1540 etiologies, usual, 290 Finegoldia magna, 911 Fusarium, 2058, 2067, 2068 Gram-positive anaerobic cocci (GPAC), 910–911 Haemophilus influenzae, 669 human immunodeficiency virus-related, 1440 human metapneumovirus, 1509 influenza virus, 1471 Legionella pneumophila, 890 measles, 1520 MERS-CoV, 1569 Microascus trigonosporus, 2075 Micrococcaceae, 361 microsporidia, 2210 Mycoplasma, 1091–1092 Neisseria meningitidis, 637 Nocardia cyriacigeorgica, 516 Nocardia otitidiscaviarum, 517 Orientia, 1124 parainfluenza virus, 1488 Pasteurella, 655 phaeohyphomycoses, 2163 Pneumocystis carinii, 2015–2026 Pneumocystis jirovecii, 1440, 1978 Prevotella, 973 Pseudomonas, 776 Pseudomonas aeruginosa, 774–775 respiratory syncytial virus (RSV), 1500 Rickettsia, 1124 Rothia mucilaginosa, 361 Scopulariopsis brevicaulis, 2075 Staphylococcus, 357, 360 Stenotrophomonas maltophilia, 794 Streptococcus mitis group, 386 Talaromyces marneffei, 2046 Trichomonas vaginalis, 2414 varicella-zoster virus, 1705 Vibrio metschnikovii, 766

viruses, specimens and methods for detection of, 1408 Pneumonic plague, 125, 742 Pneumonitis arenaviruses, 1673 Ascaris lumbricoides, 2451 enteric adenoviruses, 1622 herpes simplex virus (HSV), 1689 human herpesvirus 6 (HHV-6), 1756 human T-cell lymphotropic viruses (HTLVs), 1461 microsporidia, 2210, 2213 Toxoplasma gondii, 2375 Pneumonitis, hypersensitivity nitrofurantoin, 1196 Saccharomonospora, 520 Saccharopolyspora, 520 Thermoactinomyces, 520 Pneumovirinae (subfamily), 1398, 1487, 1498, 1508 Pneumovirus (genus), 1398, 1498, 1508 Pneumoviruses avian, 1508 human, 1508–1512 p-Nitrophenyl-β-D-glucopyranoside, 318 POCkit-HSV-2 test, 1694 Point-of-care tests, 45, 178 Chlamydia trachomatis, 1113 Epstein-Barr virus, 1743 human immunodeficiency virus, 1445 pol gene/protein, HIV, 1436–1437, 1441 polA gene, Treponema pallidum, 1063 Poliomyelitis, 1538–1540 laboratory tests suggested for, 125 specimen selection, 1541 Poliovirus, 1536 clinical significance, 1538–1540 genome organization, 1537 lyophilization of, 167 serotypes, 1537 structure of particle, 1538 transmission, 2513 typing systems, 1544–1545 vaccine, 148, 1539–1540, 1544–1545, 1804 wild, 1538–1539 Polyarthritis, mumps virus and, 1493 Polycytella hominis, 2180 Polyene(s), 2228–2229 Polyene resistance, 2237, 2239, 2243 epidemiology, 2243 intrinsic, 2237 mechanisms, 2243 Polyhexamethyl biguanide, for Acanthamoeba, 2394 Polymerase chain reaction, see PCR Polymerase inhibitors, hepatitis C virus, 1879–1880 Polymyositis, HTLVs and, 1461 Polymyxin(s), 1192–1193 adverse effects, 1193 concentration in serum, 1199 mechanism of action, 1192 pharmacology, 1192–1193 spectrum of activity, 1193 Polymyxin B, 1192–1193, 1199 Polymyxin B-lysozyme-EDTA-thallous acetate agar, 342 Polymyxin resistance, 1232 Polyomaviridae (family), 1803 taxonomic classification, 1398, 1400 virion morphology, 1401 Polyomavirus (genus), 1398, 1803 Polyomavirus-associated nephropathy (PVAN), 1804–1807, 1811–1812

SUBJECT INDEX Polyomaviruses, 1803–1812 antigen detection, 1806–1807 antiviral susceptibilities, 1811 clinical significance, 1804–1806 collection, transport, and storage of specimens, 1806 cytopathic effect (CPE), 1810 description of agents, 1803–1804 detection and identification methods, 1435 direct examination, 1806–1810 epidemiology and transmission, 1804 evaluation, interpretation, and reporting of results, 1811–1812 in situ hybridization (ISH), 1807–1808 isolation and culture procedures, 1810 microscopy, 1806–1808 nucleic acid amplification tests (NAATs), 1807, 1809–1810 commercial products, 1810 internal controls, 1809 methods, 1809–1810 positive controls and standards, 1809 template extraction, 1807, 1809 nucleic acid detection, 1807, 1809–1810 in situ hybridization (ISH), 1807–1808 Southern blotting, 1807 serologic tests, 1810 taxonomy, 1803 Polyphasic identification of fungi, 1940 Polyphasic species concept, 256 Polyvinyl alcohol (PVA) parasitology preservatives/fixatives, 2310– 2312 stool specimen preservation, 2303–2304 Polyvinylpyrrolidone (PVP) iodine, 185–186 Pontiac fever, 890 Porcine reproductive and respiratory syndrome (PRRS), 1673 Pork tapeworm, see Taenia solium Porocephalus crotali, 2516 Porphyrin test, 675–676 Porphyromonadaceae (family), 652–653, 967 Porphyromonas antibacterial resistance patterns, 1347 antimicrobial susceptibilities, 1172, 1174– 1175, 1177–1178, 1180, 1183– 1184, 1190, 1347 β-lactamase, 1346–1347 characteristics of genus, 970–971 clinical significance, 971 identification, 976, 978–979 isolation procedures, 976 taxonomy and description of genus, 967– 969 Porphyromonas asaccharolytica, 967, 971, 978– 979 Porphyromonas bennonis, 967, 971, 979 Porphyromonas cangingivalis, 971 Porphyromonas canoris, 971 Porphyromonas cansulci, 971 Porphyromonas catoniae, 967, 971, 978–979 Porphyromonas endodontalis, 967, 971, 978– 979 Porphyromonas gingivalis, 229, 967, 971, 979, 1057 Porphyromonas macacae, 971 Porphyromonas somerae, 967, 971, 979 Porphyromonas uenonis, 967, 971, 978–979 Port-A-Cul transport system, 975 Portal pyemia, Aeromonas, 754 Pos ID family, 364 Posaconazole, 2226 antifungal susceptibility testing, 2255– 2273

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 127

Aspergillus, 2044–2045 Candida, 2004–2005 chromoblastomycosis, 2167 dermatophytes, 2145 dimorphic fungi, 2121–2122 eumycotic mycetoma fungi, 2181–2182 Fusarium, 2069 hyaline fungi, 2077 Leishmania, 2361 melanized fungi, 2167 mucormycosis, 2097 phaeohyphomycosis, 2167 scedosporiosis, 2167 spectrum of activity, 2224, 2226 sporotrichosis, 2167 Talaromyces marneffei, 2048 yeast species, MICs for, 2005 Posaconazole resistance, 2226, 2239 Positive control, 80 Positive predictive value, 92–93 Postexposure management, 178 Postherpetic neuralgia, herpes zoster and, 1705–1706 Post-HSCT acute limbic encephalitis (PALE), 1756 Post-kala-azar dermal leishmaniasis, 2358– 2362 Postpartum sepsis, Gardnerella vaginalis and, 479 Posttransplant lymphoproliferative disorders (PTLD), Epstein-Barr virus, 1740– 1741, 1746–1747 Potassium hydroxide (KOH), 1956, 2137, 2146–2148 fungi, 1970 yeasts, 1995 Potassium hydroxide (10%) with lactophenol cotton blue (LPCB), 1956 Potassium iodide, for entomophthoromycosis, 2099 Potassium tellurite agar, 342 Potato dextrose agar, 1952, 1961, 2068 Potato flake agar, 1952, 1961 Powassan fever, 2523 Powassan virus, 1645 Poxviridae (family), 1828 taxonomic classification, 1395, 1397, 1398, 1400, 1402 virion morphology, 1401 Poxviruses, 1828–1837 antigen detection, 1833 clinical significance, 1830–1832 antiviral therapy, 1831–1832 molluscipoxvirus, 1831 orthopoxviruses, 1830–1831 parapoxviruses, 1831 yatapoxviruses, 1831 description of agents, 1828 detection and identification methods, 1435 diagnostic tests, 1832 direct detection, 1832–1835 epidemiology and transmission, 1828–1830 molluscipoxvirus, 1829–1830 orthopoxviruses, 1829 parapoxviruses, 1829–1830 yatapoxviruses, 1829 evaluation, interpretation, and reporting of results, 1837 genomes, 1828 identification, 1835 isolation, 1835 microscopy, 1832–1833 nucleic acid detection, 1833–1835

n cxxvii

serologic tests, 1835–1837 specimen collection and handling, 1406, 1832 taxonomy, 1828–1829 virion morphology, 1828 PPLO agar, 342 PPLO broth base without crystal violet, 340–341 Pragia fontium, 718, 727 Pramoxine, 2515 PRAS media anaerobic Gram-negative rods, 975 Clostridium identification, 954 Praziquantel, 2532–2533 adverse effects, 2533 Diphyllobothrium latum, 2473 Dipylidium caninum, 2501 Echinococcus granulosus, 2476 Fasciola, 2490 Fasciolopsis buski, 2490 heterophyid trematodes, 2490 indications for, 2533 mechanism of action, 2532 nanophyetiasis, 2490 Paragonimus, 2487 pharmacokinetics, 2532–2533 schistosomes, 2486, 2564, 2567 spectrum of activity, 2533 Taenia saginata, 2474 Taenia solium, 2475, 2476 Praziquantel resistance, 2551, 2555–2556, 2564, 2567 Precipitation reactions, 95–96 Precipitin curve, 95 Predictive value, of immunoassays, 92–93 Pregnancy arenaviruses, 1673, 1675 cytomegalovirus, 1718–1719 hepatitis E virus, 1589 human immunodeficiency virus transmission, 1438 parvovirus B19, 1820 polyomaviruses, 1804 Toxoplasma gondii and, 2373–2375, 2379– 2380 Trypanosoma cruzi, 2363 varicella-zoster virus infection, 1704–1706 Premier Adenoclone, 1624 Premier Adenoclone–Types 40/41, 1624– 1625 Premier Campy Campylobacter assay, 1002 Premier Coccidioides EIA, 1971, 2121 Premier Cryptococcal Antigen, 1997 Premier Platinum HpSA, 1019 Premier Platinum HpSA PLUS, 1019 Premier Rotaclone, 1624 Prenatal screening, for rubella virus, 1530 Prescottella, 518 Preservation, see also Storage of microorganisms for parasitology, 2310–2312 stool specimens for parasitology, 2300– 2304 formalin, 2301–2303 modified polyvinyl alcohol, 2303–2304 polyvinyl alcohol (PVA), 2303–2304 Schaudinn’s fluid, 2303–2304 single-vial collection systems, 2303– 2304 sodium acetate-acetic acid-formalin (SAF), 2302–2303 storage of microorganisms freeze-drying (lyophilization), 164–165 long-term preservation methods, 162– 165

cxxviii n

SUBJECT INDEX

Preservation, see also Storage of microorganisms (continued) short-term preservation methods, 161– 162 ultralow-temperature freezing, 162–164 Preservatives, 320–321 PreservCyt, 1414 Presurgical skin disinfection, 187–188 PreTect HPV-Proofer, 1790, 1793 PREVI Isola, 49 Prevotella antibacterial resistance patterns, 1347 antimicrobial susceptibilities, 983–984, 1172, 1174, 1175, 1177–1178, 1180, 1183–1185, 1187, 1190, 1347, 1351 β-lactamase, 1342, 1346–1347 characteristics of genus, 970–971 clinical significance, 972–973 direct examination, 975 epidemiology and transmission, 969 identification, 976–977, 979–980, 982 isolation procedures, 976 taxonomy and description of genus, 967– 969 Prevotella amnii, 973, 980 Prevotella aurantiaca, 979–980 Prevotella baroniae, 972, 980 Prevotella bergensis, 973, 980 Prevotella bivia, 973, 980, 1347 Prevotella buccae, 972–973, 980 Prevotella buccalis, 973, 975, 980 Prevotella copri, 230, 980 Prevotella corporis, 973, 979–980 Prevotella dentalis, 980 Prevotella denticola, 972–973, 979–980 Prevotella disiens, 973, 980 Prevotella enoeca, 980 Prevotella fusca, 979–980 Prevotella heparinolytica, 967, 973, 980 Prevotella histicola, 973, 979–980 Prevotella intermedia, 972–973, 977, 979, 982 Prevotella jejuni, 979–980 Prevotella loescheii, 979–980 Prevotella maculosa, 980 Prevotella marshii, 979–980 Prevotella massiliensis, 979 Prevotella melaninogenica, 969, 972–973, 977, 979–980, 1347 Prevotella micans, 979–980 Prevotella multiformis, 980 Prevotella multisaccharivorax, 980 Prevotella nanceiensis, 980 Prevotella nigricans, 972–973, 977, 979–980, 982 Prevotella oralis, 980 Prevotella oris, 973, 980 Prevotella pallens, 977, 979–980 Prevotella ruminicola, 975 Prevotella saccharolytica, 980 Prevotella salivae, 973, 980 Prevotella shahii, 979–980 Prevotella stercorea, 980 Prevotella tannerae, 972, 980; see also Alloprevotella tannerae Prevotella timonensis, 973, 980 Prevotella veroralis, 980 Prevotella zoogleoformans, 967, 980 Prevotellaceae (family), 967 Priapism, Spanish fly and, 2521 Primaquine, 2539, 2564 adverse effects, 2539 mechanism of action, 2539 pharmacokinetics, 2539

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 128

spectrum of activity, 2539 Primaquine resistance, 2552 Primary amebic meningoencephalitis (PAM), 2387, 2389–2390, 2392, 2395 Primary effusion lymphoma (PEL), 1763– 1764 Primers, see PCR (polymerase chain reaction) Prions, 1859–1864 antigen detection, 1863–1864 classification, 1400 clinical significance, 1861 collection, transport, and storage of specimens, 1861–1862 safety and security, 1861–1862 shipping, 1862 specimen collection, 1862 CSF analysis, 1864 description of agent, 1859 direct examination, 1862–1864 epidemiology and transmission, 1859–1861 evaluation, interpretation, and reporting of results, 1864 laboratory-acquired infections, 177–178 microscopy, 1862–1863 mutations in PRNP, 1862 nucleic acid detection, 1863–1864 taxonomy, 1400, 1859 Prism HBc assay, 1848 Prism HBsAg, 1847 Prism series of sequence detection systems, 74 Pro HMPV assay, 1506, 1511 Pro HMPV Real Time assay, 1510 Probenecid, 1171 ProbeTec herpes simplex virus (HSV 1 & 2) QX amplified DNA assays, 1691 ProbeTec strand displacement amplification (SDA), 1110–1111 Procercoid, 2471 Procerovum, 2482 Process improvement, as clinical microbiology laboratory, 47 Proctitis Campylobacter, 1001 Chlamydia trachomatis, 1108 herpes simplex virus (HSV), 1689 viruses, specimens and methods for detection of, 1408 Proctocolitis, Helicobacter, 1017–1018 Prodesse ProAdeno+ assay, 1775 Prodesse ProGastro SSCS, 1002 Prodesse ProParaflu+, 1490 Product contamination, outbreak investigations on, 126–128 ProFAST+ assay, 1477 ProFlu+, 1477, 1506 ProGastro SSCS, 690 Proglottids, 2324, 2471–2476 Progressive multifocal leukoencephalopathy (PML), 1407, 1756, 1804–1806, 1808, 1811–1812 ProMED-mail, 128 Promicromonospora, 477 PROMPT system, 33, 1296 Propamidine isethionate, for Acanthamoeba, 2394 Propionibacterium, 474 antimicrobial susceptibilities, 931, 1183, 1187, 1189, 1194, 1348 biochemical characteristics, 929 blood culture contaminant, 18 clinical significance, 922, 924

identification, 438, 927, 930 in skin microbiome, 232 taxonomy and description, 920–921 Propionibacterium acidifaciens, 924, 929 Propionibacterium acnes, 922, 924, 926–927, 929–930 antimicrobial susceptibilities, 1175, 1189– 1190, 1197, 1351 clinical significance, 905 Propionibacterium avidum, 494, 929 Propionibacterium granulosum, 929 Propionibacterium propionicum, 924, 929 Propioniferax, 474, 921, 929 Propioniferax innocua, 920, 929 Propionimicrobium, 921, 929 Propionimicrobium lymphophilum, 920, 929 ProSpecT Campylobacter assay, 1002 ProSpecT Cryptosporidium, 2441 ProSpecT Cryptosporium microplate assay, 2295 ProSpecT Entamoeba histolytica, 2295 ProSpecT for gastroenteritis viruses, 1624 ProSpecT Giardia, 2295 ProSpecT Giardia EZ, 2295, 2411 ProSpecT Giardia/Cryptosporidium, 2295, 2441 Prostate fluid, for fungi, 1944, 1946–1947, 1950, 1953 Prostate specimen collection, transport, and storage guidelines, 278 Prostatic abscess, Burkholderia, 794 Prostatitis Campylobacter, 1000 etiologies, usual, 291 Prosthesis-associated infection anaerobic Gram-negative rods, 972 Bacillus licheniformis, 442 Coniochaeta mutabilis, 2076 Coprinus cinereus, 2075 Finegoldia magna, 911 Helcococcus, 425 Lactococcus, 424 Mycobacterium, 599, 600 Paenibacillus alvei, 443 Parvimonas micra, 911 Pasteurella, 655 Propionibacterium acnes, 905, 924 Scopulariopsis brevicaulis, 2075 Staphylococcus, 360 Tannerella forsythia, 972 Veillonella, 911 Prosthetic devices, collection of specimens from, 281 Prostration, variola virus and, 1830 Protease inhibitor(s) hepatitis C virus, 1879 human immunodeficiency virus (HIV), 1440, 1871, 1874–1877 Protease inhibitor resistance hepatitis C virus, 1608, 1901–1902 human immunodeficiency virus (HIV), 1897–1898 Proteinase K-resistant prion protein, 1859 ProteinChip array, 263 Proteobacteria (phylum), 232, 967–969 Proteomics antifungal susceptibility testing, 2267 identification systems, 35–39 Nocardia, 525–526 Proteose peptone-yeast extract-glucose (PPYG) medium, 2315 Proteromonas lacertae, 2406 Proteus antibiotic resistance, 1227, 1232, 1234

SUBJECT INDEX antimicrobial susceptibilities, 727–730, 1174, 1177–1178, 1186–1187, 1193, 1195 antimicrobial susceptibility testing, 1266 description of genus, 715 epidemiology, transmission, and clinical significance, 720–721 identification, 724–727 swarming, 722 Proteus hauseri, 717, 724 Proteus mirabilis, 717, 720, 724–725, 727– 729, 1178 antibiotic resistance, 1227, 1232 β-lactamases, 1299 Proteus myxofaciens, 717 Proteus OX-19 agglutinating antibodies, 1128–1130 Proteus penneri, 717, 720, 724, 730 Proteus vulgaris, 199, 717, 720, 724–726, 730 Proto-fix CLR, 2311 Protoparvovirus (genus), 1818 Prototheca, 2004 Prototheca wickerhamii, 1989–1990, 2004, 2116 Protozoa, see also specific organisms amebae, 2399–2408 antiprotozoal agents, 2530, 2535–2545 ciliates, 2400, 2416–2417 coccidia, 2425–2431 commercial kits for immunodetection of serum antibodies, 2296 flagellates, 2400, 2408–2416 storage methods, 166 taxonomy and classification, 2285–2288 Protozoa (kingdom), 1938–1939 Protozoa (phylum), 2387 Prove-it Sepsis microarray assay, 1997 Providencia antibiotic resistance, 1232 antimicrobial susceptibilities, 1178, 1186– 1187, 1193 description of genus, 715 epidemiology, transmission, and clinical significance, 720–721 identification, 724–726 Providencia alcalifaciens, 717, 720–721, 724 Providencia burhodogranariea, 717 Providencia heimbachae, 717, 724, 730 Providencia rettgeri, 717, 721, 724, 730 Providencia rustigianii, 717, 724 Providencia sneebia, 717 Providencia stuartii, 717, 720, 724, 729–730, 1217 Providencia vermicola, 717 Prozone effect, 1977 PRPN gene, 1859, 1862–1864 Pruritus hypersensitivity reactions, 2515 illusory parasitosis, 2521 Mansonella, 2468 Onchocerca volvulus, 2466 quinolones, 1180 rifaximin, 1195 scabies, 2516 PSEA, hepatitis C virus, 1607 Pseudallescheria, 1937, 2159, 2173–2174, 2178, 2180–2181 antifungal susceptibility testing, 2271 cycloheximide inhibition, 1955 Pseudallescheria apiosperma, 2155, 2159 Pseudallescheria boydii, 1938, 2067, 2155– 2156, 2159, 2163, 2166–2167, 2173 antifungal resistance, 2243 cycloheximide inhibition, 1951

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 129

microscopy, 1967 Pseudamphistomum aethiopicum, 2481 Pseudamphistomum truncatum, 2481 Pseudoallescheriosis, specimens for, 1947 Pseudochrobactrum, 824 Pseudochrobactrum asaccharolyticum, 626–627, 823–824 Pseudochrobactrum saccharolyticum, 824 Pseudocitrobacter, 715, 720 Pseudocitrobacter anthropi, 720 Pseudocitrobacter faecalis, 720 Pseudoclavibacter chemotaxonomic features, 475 description of genus, 478 identification, 495–496 Pseudoclavibacter alba, 478 Pseudoclavibacter bifida, 478 Pseudocowpox, 1828–1829, 1837 Pseudoflavonifractor, 967 Pseudoflavonifractor capillosus, 967, 970, 984 Pseudofungi, 1936 Pseudohyphae, 1935, 1939, 1942, 1966, 1969, 1973, 2153 Pseudomembranous colitis carbapenems, 1177 cephalosporins, 1175 clindamycin, 1185 Clostridium difficile, 944 macrolides, 1183 penicillins, 1173 tetracyclines, 1187 Pseudomicrodochium, 2156 Pseudomonas, 773–784, 791 antibiotic resistance, 1217 antimicrobial resistance, 781–782 mechanisms, 781–782 multidrug, 782 antimicrobial susceptibilities, 782–784, 1172–1175, 1177–1178, 1181, 1193, 1195–1196 carbapenemases, 1300 classification in genus, 264 clinical significance, 774–776 collection, transport, and storage of specimens, 776 commercial sources of chromogenic agar media for, 326 description of agent, 773–774 direct examination, 776 MALDI-TOF MS, 776 microscopy, 776 nucleic acid detection, 776 epidemiology and transmission, 774 evaluation, interpretation, and reporting of results, 784 identification, 777–780 characteristics, 779 commercial systems, 778, 780 fluorescent group, 777–778 nonfluorescent group, 778 isolation procedures, 722, 776–777 serologic tests, 780 taxonomy, 773 typing systems, 780 Pseudomonas aeruginosa, 614–615, 895 acetamide agar for, 327 antibiotic resistance, 1214, 1218–1220, 1224–1228, 1232, 1234 antimicrobial resistance, 781–782 antimicrobial susceptibilities, 782, 1173– 1177, 1179–1181, 1184, 1186– 1187, 1191–1193, 1195 antimicrobial susceptibility testing, 1254– 1256, 1258–1260, 1266–1267, 1279–1280

n cxxix

as ESKAPE pathogen, 714 biochemical characteristics, 624–629 blood culture, 18 carbapenemases, 781–782, 1300–1301 clinical significance, 774–775 direct examination, 776 disinfection, 194–195 endoscope contamination outbreak, 199 epidemiology and transmission, 774 evaluation, interpretation, and reporting of results, 784 identification, 777–780 characteristics, 779 commercial systems, 778, 780 in cystic fibrosis patients, 299 isolation procedures, 777 reference strain, 1264, 1267 serologic tests, 780 taxonomy, 773 typing systems, 780 Pseudomonas alcaligenes, 615, 628–629, 773, 775–776, 778 Pseudomonas aureofaciens, see Pseudomonas chlororaphis Pseudomonas chlororaphis, 773 Pseudomonas extorquens, see Methylobacterium mesophilicum Pseudomonas fluorescens, 232, 615 antibiotic resistance, 1226 antimicrobial susceptibilities, 784 as protective skin organism, 232 biochemical characteristics, 624–629 clinical significance, 775–776 epidemiology and transmission, 774 identification, 778–780 characteristics, 779 commercial systems, 780 isolation procedures, 777 mupirocin production, 1197 taxonomy, 773 Pseudomonas isolation agar base with glycerol, 342 Pseudomonas luteola, 615, 626–627, 773–776, 778 Pseudomonas maltophilia, 792 Pseudomonas marginalis, 773 Pseudomonas mendocina, 628–629, 773, 775– 776, 778 Pseudomonas mesophilica, see Methylobacterium mesophilicum Pseudomonas monteilii, 624–625, 773, 775– 776, 778 Pseudomonas mosselii, 624–625 Pseudomonas oryzihabitans, 615, 626–627, 773–779 Pseudomonas paucimobilis, see Sphingomonas Pseudomonas pseudoalcaligenes, 615, 623, 628– 629, 773, 775–776, 778 Pseudomonas putida antibiotic resistance, 1226 antimicrobial susceptibilities, 784 biochemical characteristics, 624–625, 628– 629 clinical significance, 775–776 identification, 778–780 characteristics, 779 commercial systems, 780 isolation procedures, 777 taxonomy, 773 Pseudomonas stutzeri, 196, 615, 628–629, 800, 1177 antibiotic resistance, 1226 clinical significance, 775–776

cxxx

n

SUBJECT INDEX

Pseudomonas stutzeri (continued) identification, 778–779 taxonomy, 773 Pseudomonas syringae, 774 Pseudomonas veronii, 624–625, 773, 775–776, 778 Pseudomonic acid A, 1197 Pseudonocardia chemotaxonomic and lysosome growth characteristics, 509 description of genus, 508 identification, 523 morphologic characteristics, 508 taxonomy, 505 Pseudo-outbreak, 112 Pseudophyllidea (order), 2291, 2471, 2502 Pseudoramibacter, 921 Pseudoramibacter alactolyticus, 922, 924, 926– 927, 930 Pseudoterranova, 2495 Pseudoterranova decipiens, 2493 Pseudozyma, 2001 clinical significance, 1994 description of agents, 1990–1991 taxonomy, 1985 Pseudozyma alboarmenica, 1994 Pseudozyma antarctica, 1985, 1994 Pseudozyma aphidis, 1985, 1991, 1994 Pseudozyma crassa, 1994 Pseudozyma paraantarctica, 1985, 1994 Pseudozyma siamensis, 1994 Pseudozyma thailandica, 1985, 1994 Psittacosis, 223, 1109 PSMEA, hepatitis C virus, 1607 Psocoptera (order), 2510 PSORTb, 233 Psychrobacter, 813, 822 Psychrobacter faecalis, 632–633, 820, 822 Psychrobacter immobilis, 822 Psychrobacter phenylpyruvica, 820–822 Psychrobacter phenylpyruvicus, 632–633 Psychrobacter pulmonis, 632–633, 820, 822 Psychrobacter sanguinis, 632–633, 820–822 Pthirus pubis, 2330 Pubic louse, 2330, 2510–2511 Public Health Security and Bioterrorism Preparedness and Response Act of 2002, 167 Puerperal sepsis, 385 Pulex, 2507 Pulex irritans, 2509 Pulmonary disease/infection, see also Lung disease/infection Actinomyces, 923 adiaspiromycosis, 2115 Blastomyces dermatitidis, 2114 Brucella, 865 Campylobacter, 1001 Chlamydia pneumoniae, 1109 dirofilariasis, 2499 hantaviruses, 1662 Histoplasma capsulatum, 2114 human metapneumovirus, 1509 hyaline fungi, 2075–2076 mucormycosis, 2089 Mycobacterium, 538, 542–544, 599 Nocardia, 515, 518 Paracoccidioides brasiliensis, 2115 Paragonimus, 2487 parainfluenza virus, 1488 phaeohyphomycoses, 2163 Pseudozyma, 1994 Rhodococcus equi, 518–519 sporotrichosis, 2164

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 130

Strongyloides stercoralis, 2457 Tropheryma whipplei, 1161 Tsukamurella paurometabola, 519 Veillonella, 911 Pulmonary hemorrhage, Leptospira, 1030 Pulsed-field gel electrophoresis (PFGE) Brachyspira, 1065 Burkholderia, 803 Campylobacter, 1006 Clostridium, 956 described, 136 DNA fingerprint analyzed by, 128 Enterobacteriaceae, 726 Enterococcus, 412–413 Escherichia coli, 695 forensic molecular epidemiology use of, 148 Haemophilus, 677 in RFLP analysis, 136 molecular surveillance use of, 146 nontuberculous mycobacteria (NTM), rapidly growing, 605 nontuberculous mycobacteria (NTM), slowly growing, 585 Pseudomonas, 780 PulseNet and, 146, 151 Salmonella, 704 Staphylococcus, 367–368 Streptococcus, 395–396 Ureaplasma, 1097 Vibrionaceae, 769 yeasts, 2004 Yersinia, 747 PulseNet, 128, 141, 146, 151 Pulvinar sign, 1861 Punctate, 1942 Puritan universal transport medium, 1410 Purpureocillium, 2064, 2073, 2076 Purpureocillium lilacinum, 2064, 2067, 2075, 2076 antifungal resistance, 2243 antifungal susceptibility testing, 2271 Pus specimen, for fungi, 1945, 1947–1948 Puumala virus, 1660–1661 PVA-containing preservatives and fixatives, 2312 PVAN, see Polyomavirus-associated nephropathy PVL Evigene kit, 366 Pycnidia, 1939, 1942, 2058 Pycnidiospore, 1942 Pycnothyriales, 1939 Pyelonephritis etiologies, usual, 291 hyaline fungi, 2076 Mycoplasma, 1092 Pygidiopis, 2482 PyloriTek, 1019 Pyogenic liver abscess, Klebsiella pneumoniae, 718, 723 Pyothorax, Nocardia otitidiscaviarum, 517 PYR test, 319, 350–352, 391 Pyramidobacter piscolens, 968, 974, 980–981 Pyrantel pamoate, 2535 adverse effects, 2535 Enterobius vermicularis, 2454 mechanism of action, 2535 pharmacokinetics, 2535 spectrum of activity, 2535 Pyrazinamidase, 1359 Pyrazinamide activity, 1359 adverse effects, 1359 antimicrobial susceptibility testing, 1363, 1365–1367

for Mycobacterium infection, 1358–1359 Pyrazinamide resistance, 1357–1359, 1366– 1367 Pyrenochaeta, 2174 Pyrenochaeta mackinnonii, 2173–2174 Pyrenochaeta nobilis, 2174 Pyrenochaeta romeroi, 2174 Pyriform, 1942 Pyrimethamine, 1185, 1191, 2540–2541 Cystoisospora belli, 2430–2431 Toxoplasma gondii, 2381–2382 Pyrimethamine resistance, 2551–2553 Pyrimethamine-sulfadoxine, 2540–2541 Pyroglyphid mites, 2517 Pyronaridine, 2564 Pyronaridine resistance, 2552 Pyrosequencing, 70, 76 anaerobic bacteria, 907 antiviral susceptibility testing, 1916–1918 classification and identification of bacteria and, 261 Mycobacterium, 581, 604 Nocardia, 525 yeast identification, 2003 Pyrrolidonyl aminopeptidase activity, 319, 616 Pyrrolidonyl arylamidase (PYR), 319, 350– 352, 391 Pythiaceae (family), 2200 Pythiales (order), 1939, 2200 Pythiosis, 2200–2203 Pythium insidiosum, 1936, 1939, 2199–2203 antimicrobial susceptibility, 2203 clinical significance, 2198, 2201 collection, transport, and storage of specimens, 2201 description, 2200 direct examination, 2201–2202 epidemiology and transmission, 2198, 2200–2201 evaluation, interpretation, and reporting of results, 2203 identification, 2203 isolation, 2202–2203 microscopy, 2200–2202 nucleic acid detection, 2202 phylogeny, 2197–2198 serologic tests, 2203 taxonomy, 2198, 2199–2200 Q fever, 222–223, 1150–1155 acute, 1152 antimicrobial susceptibilities, treatment, and prevention, 1155 biothreat agent, 222–223 chronic, 1152 clinical significance, 1151–1153 collection, transport, and storage of specimens, 1153 description of agent, 1150 epidemiology and transmission, 222–223, 1150–1151 evaluation, interpretation, and reporting of results, 1155 in children, 1153 in pregnancy, 1152–1153 pathogenesis, 1151–1152 post-Q fever fatigue syndrome, 1153 serologic tests, 1154–1155 vaccine, 1155 QBC microhematocrit centrifugation, 2336 Q-FAME, 34 QIAcube, 1542 QIAsymphony system, 74, 1542

SUBJECT INDEX QIME (Quantitative Insights into Microbial Ecology), 246 Qinghaosu, 2539–2540 Quadriplegia, in tick paralysis, 2516 Quality control antifungal susceptibility testing, 2262, 2270 antimicrobial susceptibility testing dilution methods, 1263–1264 batch and lot QC, 1264 frequency, 1264 MIC ranges, 1264 reference strains, 1263–1264 antimicrobial susceptibility testing disk diffusion method, 1267 frequency of testing, 1267 reference strains, 1267 special disk tests, 1267 zone-of-inhibition diameter ranges, 1267 antimicrobial susceptibility testing of anaerobes, 1343–1345 automated laboratory system, 52 for molecular methods, 79–80 laboratory, 45 phenotypic methods for detecting antibacterial resistance, 1286, 1288–1289, 1291, 1295–1298, 1303 storage of microorganisms, 162 validation of subtyping methods, 149 Quality Control for Molecular Diagnostics (QCMD) EQA panel, 1577 Quality Control for Molecular Diagnostics proficiency program, 80 Quambalaria, 2062 Quambalaria cyanescens, 2062, 2071 Quambalariaceae (family), 2071 QuantiFERON assay, for cytomegalovirus, 1722 QuantiFERON gold in-tube (QFNG-IT) assay, 555–556, 576 QuantiFERON-CMV, 1730 Quantitative culture, 292 of urine, 304 Quantitative molecular assays, 73, 81–82 Quantitative PCR coccidia, 2430 herpes simplex virus (HSV), 1696 human bocavirus, 1824 human herpesvirus 6 (HHV-6), 1759– 1760 parvovirus B19, 1819, 1821–1823 Quantitative real-time PCR enteric adenoviruses, 1628 Epstein-Barr virus, 1742, 1744 human herpesvirus 6 (HHV-6), 1759 human metapneumovirus, 1511 Quantum dots, 10 Quaternary ammonium compounds (quats), 195 Queensland tick typhus, 1125 Quenching, 11 Quick Chek Giardia duodenalis, 2412 QuickFISH, 1997 QuickVue Influenza A+B test, 1474 QuickVue Influenza test, 1474 QuickVue RSV, 1502, 1504 Quidel Molecular Influenza A+B, 1477 Quidel Molecular RSV+HMPV assay, 1511 Quinacrine, for Giardia duodenalis, 2412 Quinidine, 2537–2538 adverse effects, 2538 mechanism of action, 2537

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 131

pharmacokinetics, 2537–2538 spectrum of activity, 2538 Quinine, 2349, 2537–2538, 2564 adverse effects, 2538 mechanism of action, 2537 pharmacokinetics, 2537–2538 spectrum of activity, 2538 Quinine resistance, 2552 Quinoline derivatives, 2536–2539 4-aminoquinolines, 2536–2537 8-aminoquinolines, 2539 cinchona alkaloids, 2537–2538 synthetic compounds, 2538–2539 Quinolone(s), 1178–1180 adverse effects, 1180, 1361 antimicrobial susceptibility testing, 1256, 1260 for Mycobacterium infection, 1361 mechanism of action, 1179 pharmacology, 1179 spectrum of activity, 1179–1180 Quinolone resistance, 1179–1180, 1232– 1233 alterations in target enzymes, 1232–1233 common associations of resistance mechanisms, 1215 due to decreased intracellular accumulation, 1233 Mycobacterium tuberculosis complex, 361, 1357–1358, 1367 Pseudomonas aeruginosa, 782 Quintan fever, 876 Quinupristin-dalfopristin, 1189–1190, 1199 Quinupristin-dalfopristin resistance, 1231 QVAX, 1155 Rabbit infectivity testing (RIT), 1055, 1064, 1073 Rabbit plasma, 317 Rabies, 1633–1641 diagnosis, 1636–1637 encephalitic form, 1635 epidemiology and transmission, 1633, 1635 laboratory tests suggested for, 125 paralytic form, 1635 postexposure prophylaxis, 1635–1636 preexposure prophylaxis, 1636 treatment, 1635–1636, 1641 vaccine, 1635–1636 Rabies immune globulin (RIG), 1635 Rabies virus, 1633–1641 antigen detection, 1638–1640 antigenic typing, 1640–1641 antiviral susceptibilities, 1641 clinical significance, 1635–1636 description of agent, 1633, 1635 detection and identification methods, 1435 direct detection methods, 1636–1640 direct fluorescent antibody (DFA) test, 1637–1640 direct rapid immunohistochemistry test (DRIT), 1637–1639 disinfectant susceptibility, 1635 epidemiology and transmission, 1633, 1635 evaluation, interpretation, and reporting of results, 1641 formalin-fixed tissue, tests on, 1638–1639 identification, 1640 immunohistochemistry (IHC), 1637–1639 indirect fluorescent antibody (IFA) test, 1640–1641

n cxxxi

isolation, 1640 microscopy, 1638 neutralization tests, 1641 nuchal (neck) biopsy specimens, tests on, 1639–1640 nucleic acid detection, 1637, 1640 nucleotide sequence analysis, 1641 serologic tests, 1641 specimen collection and handling, 1407, 1636 taxonomy, 1633–1634 typing systems, 1640–1641 Radial immunodiffusion, 95 Radiculitis Epstein-Barr virus, 1739 herpes simplex virus (HSV), 1689 Radiofrequency energy, 204 Radio-frequency identification (RFID) labeling, 164 Radioimmunoassay (RIA), 91, 92, 1472 Rahnella, 722, 726 Rahnella aquatilis, 718 Raillietina, 2502 Rainbow agar O157, 342–343 Rainbow agar Salmonella, 343 Ralstonia, 791 clinical significance, 795 description of genus, 792 direct examination, 795 epidemiology and transmission, 793 identification, 797–801 typing systems, 803 Ralstonia eutropha, 791 Ralstonia insidiosa, 614, 630–631, 791, 795 Ralstonia mannitolilytica, 630–631, 791, 795 Ralstonia pickettii, 614, 615, 630–631, 791, 795 Raltegravir, 1872, 1877, 1920 Raltegravir resistance, 1447, 1897, 1899 RAMBACH agar (CHROMagar Rambach), 343 Ramichloridium mackenziei, 2161 Ramichloridium obovoideum, 2159 RAMP Flu A+B, 1474 Ramsay Hunt syndrome, 1709 Random amplification of polymorphic DNA/ arbitrarily primed PCR (RAPD/APPCR), 137 Random amplified polymorphic DNA (RAPD) analysis, 2043 Random amplified polymorphic DNA PCR (RAPD-PCR), 605 Random fragment length amplification PCR, 138–139 Raoultella description of genus, 715 epidemiology, transmission, and clinical significance, 717–719 identification, 723–725 Raoultella ornithinolytica, 717, 724 Raoultella planticola, 717–718, 723–725 Raoultella terrigena, 717–718, 723–725 RapID ANA II, 927 anaerobic Gram-negative rods, 977 Bartonella, 880 Clostridium, 954 RapID CB Plus system, 481–482, 485, 490– 491, 493 Rapid combination nontreponemal and treponemal tests, 1070 Rapid fermentation medium, 343 Rapid fluorescent focus inhibition test (RFFIT), for rabies virus, 1636–1637 Rapid HIV immunoassays, 1444–1447

cxxxii

n

SUBJECT INDEX

RapID Hp StAR, 1019 Rapid ID 32, 880 Rapid ID 32A, 954, 977, 980 Rapid influenza diagnostic test (RIDT), 1473–1476 RapID NF Plus, 778 Rapid NFT, 34 RapID NH, 641–642, 676, 678 Rapid plasma reagin (RPR), 1062, 1066, 1074 Rapid point-of-care (RPOC) tests, for hepatitis E virus, 1591–1593 RapID Strep, 428 Rapid trehalose test, 2001 Rapid treponemal tests, 1069, 1071–1072, 1074 Rapid urease test, 1019 Rapid Neg ID3, 768 Rapid-sporulation agar, 1952 Rasamsonia, 2064, 2073, 2076 Rasamsonia argillacea, 2064, 2073, 2076 Rash aminoglycosides, 1182 Anaplasma phagocyrophilum, 1139 arboviruses, 1647 arenaviruses, 1674 cephalosporins, 1175 chloramphenicol, 1193 clindamycin, 1185 Coccidioides, 2114 daptomycin, 1189 Ehrlichia chaffeensis, 1138 filoviruses, 1674 fosfomycin, 1196 hookworm, 2456 human herpesvirus 6 (HHV-6), 1754– 1756 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1762 macrolides, 1183 Mansonella, 2468 measles, 1520 monkeypox virus, 1830 monobactams, 1176 nitrofurantoin, 1196 Onchocerca volvulus, 2466–2467 Orientia, 1124 parvovirus B19, 1819, 1822 penicillins, 1173 polymyxins, 1193 quinolones, 1180 Rickettsia, 1124 rifampin, 1195 rubella, 1526 sulbactam, 1178 sulfonamides, 1192 tetracyclines, 1187 Treponema, 1061 Trypanosoma brucei, 2366 Trypanosoma cruzi, 2362 urticating caterpillar hairs, 2518–2519 varicella-zoster virus, 1709 variola virus, 1830 viruses, specimens and methods for detection of, 1406 Rat bite fever, 655 Rat tapeworm, 2502, 2507 Rat-tailed maggot, 2517, 2519 Ravn virus, 1670 Ravuconazole Fusarium, 2069 hyaline fungi, 2077 Ravuconazole resistance, 2239 Reactivation

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 132

cytomegalovirus, 1719 Epstein-Barr virus, 1739–1740, 1746 herpes B virus, 1697 herpes simplex virus (HSV), 1687–1688 human herpesvirus 6 (HHV-6), 1756 human papillomavirus, 1784 of pathogens in ticks, 2521, 2523 Toxoplasma gondii, 2374 varicella-zoster virus, 1704–1705, 1709, 1712 ReadyCells HSV system, 1692 Reagent excess assay, 92 Reagent-limited assays Reagents biochemical tests, 316–320 buffers, 320 decontamination agents, 320 dyes and indicators, 320, 321 McFarland standards, 321 mycology, 1956 parasitology, 2310–2312 preservatives, 320 virology, 1422–1423 Real de Catorce virus, 1669, 1672 RealAccurate respiratory RT-PCR kit v2.0, 1574 RealArt M. tuberculosis TM PCR reagents, 575 RealStar CMV PCR kits, 1726 Real-time PCR, 60–63 amebae, 2393 antiviral susceptibility testing, 1916, 1918 arenaviruses, 1676–1677, 1681 coronaviruses, 1571–1574, 1577 cytomegalovirus, 1725–1726 Entamoeba histolytica, 2405 filoviruses, 1676–1677, 1681 gastroenteritis viruses, 1625–1626, 1628 herpes simplex virus (HSV), 1690–1691 human herpesvirus 7 (HHV-7), 1761 human T-cell lymphotropic viruses (HTLVs), 1462 Pneumocystis, 2024 polyomaviruses, 1809–1810 poxviruses, 1833–1835 Trypanosoma cruzi, 2365 varicella-zoster virus, 1711 Real-time RT-PCR human immunodeficiency virus, 1442– 1443 human metapneumovirus, 1511 influenza viruses, 1476–1479, 1482 rhinoviruses, 1554–1555 REBA HPV-ID, 1792 Recluse spiders, 2520 Recombinant immunoblot assay (RIBA), for hepatitis C virus, 1608 recomWell Treponema IgM, 1067 Reconstitution of freeze-dried specimens, 165 Recovirus (genus), 1617 Rectal abscess, Sutterella and, 974 Rectal fistula, Bacillus pumilus and, 442–443 Rectal prolapse, Trichuris trichiura and, 2459 Rectal schistosomiasis, 2480 Rectal swab Gram stain and plating medium recommendations, 286 specimen collection, transport, and storage guidelines, 276, 301–302 viruses, collection methods and processing of specimens, 1415 Rectal tissue specimen, for parasitology, 2329, 2332 Recurrent disease, HSV, 1688

Recurrent respiratory papillomatosis, 1786 Red complex, 229 Red man syndrome, 1189 Reduviid bugs, 2508 Reference strains, for antimicrobial susceptibility testing, 1263–1264, 1267 Refractive index (index of refraction), 6 Regan-Lowe charcoal agar, 343 Regan-Lowe semisolid transport medium, 343 Regulator of viral expression (Rev) protein/ gene, HIV, 1436–1437 Regulatory issues with molecular methods, 82 Reiter’s syndrome Campylobacter, 1000 Yersinia pseudotuberculosis, 742 Rejection of specimens, 284–285 Relapsing fever, 1037–1041; see also Borrelia arthropod vectors, 2507 specimen management, 283 Renal failure Leptospira, 1030 spider envenomation, 2520 Renibacterium, 354, 361 Renispora flavissima, 2118 RENOK device, 33, 1275, 1277 Reoviridae (family), 1617–1618, 1644 taxonomic classification, 1399, 1400 virion morphology, 1401 Repetitive-sequence-based PCR (rep-PCR), 137 nontuberculous mycobacteria (NTM), rapidly growing, 605 nontuberculous mycobacteria (NTM), slowly growing, 585 Vibrionaceae, 769 Replication assay, HIV, 1443 Reporting results of molecular assays, 80–82 Reproducibility definition, 132 of subtyping method, 132 Resazurin, 2566 Resistance to disinfectants, 191, 195–196 Resolution (resolving power), 6, 7 RespiFinder 19, 1506 RespiFinder 19/19 Cy5/22, 1575 RespiFinder human metapneumovirus, 1511 RespiFinder SMART 22, 1506, 1555, 1575 RespiFinder SMART 22 Fast, 1555, 1575 Respiratory arrest, spider envenomation and, 2520 Respiratory masks, 176 Respiratory Protection Standard, 282 Respiratory syncytial virus (RSV), 1498– 1508 antigen detection, 1502–1504 antiviral susceptibilities, 1508 clinical significance, 1500–1501 cytopathic effect (CPE), 1502, 1505, 1511–1512 description of agent, 1498 detection and identification methods, 1435 DFA and IFA reagents for the detection of, 1425 direct examination, 1501–1502 direct fluorescent antibody (DFA) test, 1501–1503, 1508, 1510–1512 epidemiology and transmission, 1498–1500 evaluation, interpretation, and reporting of results, 1508 genotyping, 1505

SUBJECT INDEX identification, 1505 immunocompromised patients, 1500–1501 immunofluorescence detection in R-Mix cells, 1426 isolation procedures, 1502, 1505 laboratory tests suggested for, 125 microscopy, 1501–1502 nucleic acid amplification tests (NAATs), 1502, 1506–1508, 1511–1512 nucleic acid detection, 1502, 1506–1507 rapid antigen detection tests, 1502, 1504 rapid cell culture, 1426 serologic tests, 1505, 1508 specimen collection and handling, 1407– 1408, 1501 taxonomy, 1498 transport medium for, 1409 treatment and prevention, 1501 typing systems, 1505 virion morphology, 1498–1499 Respiratory tract microbiome of, 231 trematodes of, 2481, 2484, 2487 viruses, collection methods and processing of specimens, 1414–1415 Respiratory tract disease/infection Achromobacter xylosoxidans, 840–841, 843 adenoviruses, 1769–1773 anaerobic Gram-negative rods, 972 Anaplasma phagocyrophilum, 1139 arboviruses, 1647 Aspergillus, 2033, 2036–2037, 2044 Bacillus cereus, 443 bocavirus, 1823 Bordetella, 841 Borrelia, 1041 Chlamydia pneumoniae, 1108–1109 Citrobacter, 720 Coccidioides, 2114 cockroaches, 2513 Corynebacterium kroppenstedtii, 490 Corynebacterium minutissimum, 479 Corynebacterium pseudodiphtheriticum, 479 Corynebacterium striatum, 479 Corynebacterium ulcerans, 479 Ehrlichia chaffeensis, 1138 Eikenella corrodens, 655 enteric adenoviruses, 1618 enterovirus, 1540 etiologies, usual, 290 Finegoldia magna, 911 Haemophilus influenzae, 669 Hafnia, 721 influenza virus, 1471 Kingella, 655 monkeypox virus, 1830 Moraxella catarrhalis, 813–814, 831 mumps virus, 1493 Mycoplasma, 1091–1092, 1097–1098 Nocardia paucivorans, 516 non-spore-forming, anaerobic, Grampositive rods, 923 parechovirus, 1540–1541 Pasteurella, 655 polyomaviruses, 1804 Pseudomonas aeruginosa, 774–775 Rothia, 479 scorpion venom, 2520 Serratia, 720 specimen selection, 1541 Sporobolomyces, 1994 tick paralysis, 2516 viruses, specimens and methods for detection of, 1407–1408

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 133

Respiratory tract specimen anaerobic bacteria identification and, 905 fungi, 1946–1948, 1950 parasitology, 2294, 2299 Respiratory-V Cassette, 1555 Respirovirus (genus), 1398 Respi-Strip, 1502, 1504 ResPlex II assay, 1511, 1555 ResPlex II Panel v2.0, 1506 Reston ebolavirus, 1670 Reston virus, 1669–1670, 1672–1674, 1676, 1678, 1680 Restriction endonuclease analysis (REA), 136 adenovirus, 1776 Clostridium, 956 yeasts, 2004 Restriction endonucleases, 67 Restriction fragment length polymorphism (RFLP), 67 Aspergillus, 2043 Blastomyces dermatitidis, 2119 described, 136 Histoplasma capsulatum, 2119 influenza viruses, 1480–1481 Mycoplasma, 1097 Pseudomonas, 780 respiratory syncytial virus, 1505 Treponema pallidum, 1065 varicella-zoster virus, 1711 Retapamulin, 1197 Reticulate body, 1106–1107, 1111 Retinitis BK polyomavirus, 1805 viruses, specimens and methods for detection of, 1407 Retortamonadea (class), 2287, 2408 Retortamonadida (order), 2287 Retortamonas, 2408 Retortamonas intestinalis, 2321, 2400, 2408– 2410, 2416 Retroviridae (family), 1436, 1458 taxonomic classification, 1399, 1400 virion morphology, 1401 Reveal G3 Rapid HIV-1 antibody test, 1445 Reverse hybridization tests, for hepatitis C virus, 1605, 1607 Reverse transcriptase (RT), HIV, 1437 Reverse transcriptase activity of HBV polymerase, 1899–1900 Reverse transcriptase inhibitor(s) nonnucleoside reverse transcriptase inhibitors (NNRTIs), 1870, 1873– 1874 resistance, 1897–1898 nucleoside/nucleotide reverse transcriptase inhibitors (NRTIs), 1869–1870, 1872–1873 resistance, 1896–1898 Reverse transcriptase PCR, see RT-PCR Reverse transcriptase quantitative PCR (RTqPCR), measles virus, 1522, 1525 Reverse transcription-loop mediated amplification (RT-LAMP), influenza viruses, 1476 Reversed passive latex agglutination (RPLA), for Clostridium perfringens, 948 RevolutioN, 1099 Reye’s syndrome, 1471 RFLP, see Restriction fragment length polymorphism Rhabditida (order), 2289 Rhabditoidea (superfamily), 2289

n cxxxiii

Rhabdomyolysis, influenza virus and, 1471 Rhabdoviridae (family), 1633, 1644 taxonomic classification, 1398, 1400 virion morphology, 1401 Rhadinovirus (genus), 1762 Rheumatoid factor, 96–97, 100 Rhinitis cockroaches, 2513 Schizophyllum commune, 2075 viruses, specimens and methods for detection of, 1408 Rhinocerebral mucormycosis, 1949 Rhinocladiella, 2153–2154, 2159 Rhinocladiella aquaspersa, 1967, 2154, 2158– 2159, 2164 Rhinocladiella atrovirens, 2159 Rhinocladiella basitona, 2154, 2159 Rhinocladiella mackenziei, 2154, 2158–2159, 2163 Rhinocladiella similis, 2154, 2159 Rhinorrhea coronaviruses, 1569 human herpesvirus 6 (HHV-6), 1755 influenza virus, 1471 respiratory syncytial virus (RSV), 1500 Rhinosporidium seeberi, 2198, 2205 Rhinoscleroma, Klebsiella rhinoscleromatis, 718 Rhinosinusitis Gram-positive anaerobic cocci (GPAC), 910 Peptoniphilus, 911 Pseudomonas aeruginosa, 780 Rhinosporidiosis, 2204–2207 Rhinosporidium seeberi, 1936, 2116, 2204– 2207 antimicrobial susceptibility, 2207 clinical significance, 2198, 2205 collection, transport, and storage of specimens, 2205 description, 2205 direct examination, 2205–2206 epidemiology and transmission, 2198, 2205 evaluation, interpretation, and reporting of results, 2207 identification, 2206–2207 microscopy, 2205–2206 phylogeny, 2197–2198 serologic tests, 2207 staining, 1958 taxonomy, 2198, 2204–2205 Rhinoviruses, 1536–1537, 1551–1559 acid pH stability, 1557 antigen detection, 1553–1554 antiviral susceptibilities, 1558 cell culture, 1556 clinical significance, 1553 cytopathic effect (CPE), 1430, 1556–1557 description of agent, 1551–1552 detection and identification methods, 1435 diagnostic methods, 1558 direct examination, 1553–1555 epidemiology and transmission, 1552–1553 evaluation, interpretation, and reporting of results, 1558–1559 genome structure, 1551–1552 genotyping, 1557 identification, 1557 isolation procedures, 1556 nucleic acid detection, 1554–1555 organ culture, 1556 RT-PCR, 1552, 1554–1555, 1557–1559 serologic tests, 1557–1558

cxxxiv

n

SUBJECT INDEX

Rhinoviruses (continued) specimen collection and handling, 1407– 1408, 1553 taxonomy, 1551 temperature sensitivity, 1557 typing systems, 1557 Rhipicephalus, 2507, 2514 Rhipicephalus sanguineus, 2513, 2515 Rhizobium, 615 Rhizobium radiobacter, 626–627, 823–825 Rhizoid, 1942 Rhizomucor, 1937, 2088, 2091–2094 Rhizomucor miehei, 2092 Rhizomucor pusillus, 2088, 2092, 2095 Rhizomucor variabilis var. variabilis, 2088, 2092, 2097 Rhizopodaceae (family), 2088, 2094–2096 Rhizopodea (superclass), 2387 Rhizopus, 1937, 1974, 2087–2088, 2091–2096 Rhizopus arrhizus, 2087–2088, 2094–2095, 2097, 2099–2100 Rhizopus microsporus, 1973, 2088, 2096, 2100 Rhizopus microsporus var. azygosporus, 2099 Rhizopus microsporus var. chinensis, 2096, 2099 Rhizopus microsporus var. microsporus, 2096, 2099 Rhizopus microsporus var. oligosporus, 2096, 2099 Rhizopus microsporus var. rhizopodiformis, 2096, 2099 Rhizopus oryzae, 2088, 2094, 2099 Rhizopus schipperae, 2099 Rhizopus stolonifer, 2096, 2099 Rhodnius, 2507 Rhodobacteraceae (family), 822, 824 Rhodococcus, 550 acid-fast stain, 321 chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 518–519 description of genus, 509 G+C content, 536 identification, 438, 522–526 isolation procedures, 521 microscopy, 521 morphologic characteristics, 508 taxonomy, 504–505 Rhodococcus aurantiacus, see Tsukamurella paurometabola Rhodococcus corynebacterioides, 514 Rhodococcus equi, 491, 509, 512–513, 515, 518, 521–522, 525–526 antimicrobial susceptibilities, 1184, 1190 antimicrobial susceptibility testing, 1372– 1373 Rhodococcus erythropolis, 514 Rhodococcus fascians, 514 Rhodococcus globerulus, 514 Rhodococcus gordoniae, 514 Rhodococcus rhodochrous, 514, 526 Rhodospirillaceae (family), 829 Rhodotorula, 1986, 1990, 2263 antifungal susceptibility testing, 2263 clinical significance, 1994 description of agents, 1991 identification, 1998–1999, 2001 media, 1960 specimen collection, transport, and processing, 1948 taxonomy, 1985 Rhodotorula glutinis, 1985, 1989, 1991, 1994, 2005 Rhodotorula minuta, 1985, 1994

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 134

Rhodotorula mucilaginosa, 1985, 1991, 1994, 2005 Rhodotorula rubra, 2264 Rhomboencephalitis, enterovirus, 1540 Ribavirin adenoviruses, 1777 arenaviruses, 1674, 1681 hantaviruses, 1665 hepatitis C virus, 1601–1602, 1609–1611, 1878–1879, 1901 hepatitis E virus, 1590–1591 human metapneumovirus, 1509, 1512 parainfluenza virus, 1488 rabies virus, 1641 respiratory syncytial virus, 1501 Ribavirin resistance, 1901–1902, 1917 RiboPrinter microbial characterization system, for Staphylococcus, 365 Ribosomal Database Project (RDP), 581 Ribosome modification in aminoglycoside resistance, 1220 Ribotyping Burkholderia, 803 Clostridium, 956 PCR, 137 Vibrionaceae, 769 Rice grains, dermatophyte growth on polished, 2141 Rickettsia, 1122–1130 antimicrobial susceptibilities, 1129–1130, 1184, 1187, 1193 clinical significance, 1083, 1124–1125 collection, transport, and storage of specimens, 283, 1124, 1126 description of, 1122–1123 diagnostic tests, 1086–1087 direct detection, 1126–1128 general considerations, 1126 immunologic detection, 1126–1127 molecular detection, 1127–1128 epidemiology and transmission, 1083, 1123–1124 identification, 1128 interpretation and reporting of results, 1130 isolation procedures, 1128 phylogeny, 1123 serologic tests, 1086–1087, 1128–1129 taxonomy, 1122 Rickettsia aeschlimannii, 1122 Rickettsia africae, 1122, 1125, 1127, 1129 diagnostic tests, 1086 epidemiology and clinical diseases associated with, 1083 Rickettsia akari, 1122–1123, 1125, 1127 arthropod vector, 2507, 2511 diagnostic tests, 1086 epidemiology and clinical diseases associated with, 1083 Rickettsia australis, 1122, 1125 Rickettsia bellii, 1122–1123, 1127 Rickettsia canadensis, 1123 Rickettsia conorii, 1122–1125, 1127–1130 antimicrobial susceptibilities, 1180 arthropod vector, 2507 diagnostic tests, 1086 epidemiology and clinical diseases associated with, 1083 Rickettsia felis, 1122–1125, 1127, 1129 diagnostic tests, 1086–1087 epidemiology and clinical diseases associated with, 1083 Rickettsia heilongjiangensis, 1125 Rickettsia helvetica, 1122

Rickettsia honei, 1083, 1122, 1125, 1127 Rickettsia japonica, 1122–1123, 1125, 1127 Rickettsia massiliae, 1122 Rickettsia monacensis, 1122 Rickettsia montanensis, 1122 Rickettsia parkeri, 1122, 1124–1127 diagnostic tests, 1086 epidemiology and clinical diseases associated with, 1083 Rickettsia peacockii, 1122 Rickettsia philipii, 1124 Rickettsia prowazekii, 1122–1125, 1127–1130 arthropod vector, 2507 biothreat agent, 223 diagnostic tests, 1087 epidemiology and clinical diseases associated with, 1083 transmission and disease, 223 Rickettsia rhipicephali, 1122 Rickettsia rickettsii, 1122, 1124, 1126–1127, 1129–1130, 1136 antimicrobial susceptibilities, 1180 arthropod vector, 2507 diagnostic tests, 1087 epidemiology and clinical diseases associated with, 1083 Rickettsia sibirica, 1122–1123, 1125, 1127 Rickettsia slovaca, 1122, 1125, 1127 Rickettsia typhi, 1122–1125, 1127, 1129–1130 arthropod vector, 2507 diagnostic tests, 1087 epidemiology and clinical diseases associated with, 1083 Rickettsiaceae (family), 1122 Rickettsiales (order), 873, 1135–1136 Rickettsialpox, 1124–1126, 1129, 2507, 2511 Rickettsiosis, 2511 RIDA Gene MRSA system, 361 RIDA Quick Cryptosporidium, 2441 RIDA Quick Cryptosporidium/Giardia Combi, 2441 RIDA Quick Cryptosporidium/Giardia Entamoeba Combi, 2441 RIDA Quick for gastroenteritis viruses, 1624 RIDASCREEN Adenovirus, 1624 RIDASCREEN Astrovirus, 1624 RIDASCREEN Cryptosporidium, 2441 RIDASCREEN Norovirus, 1624–1625 RIDASCREEN Rotavirus, 1624 RIDOM, 602 Rifabutin activity, 1359 adverse effects, 1359 for Mycobacterium infection, 1359 Toxoplasma gondii, 2382 Rifabutin resistance, 1359 Rifampin, 1194–1195 activity, 1357, 1359 adverse effects, 1195, 1359 antimicrobial susceptibility testing, 1256, 1261, 1365–1367 concentration in serum, 1199 for Mycobacterium infection, 1357–1359 mechanism of action, 1194 pharmacology, 1194 spectrum of activity, 1195 Rifampin resistance, 1215, 1233, 1356–1358, 1363, 1367–1368 Rifamycins, 1194–1195 adverse effects, 1195 mechanism of action, 1194 pharmacology, 1194–1195 spectrum of activity, 1195 Rifapentine, for Mycobacterium infection, 1359

SUBJECT INDEX Rifapentine resistance, 1359 Rifaximin, 1194–1195 Rift Valley Fever virus, 1645, 1647, 1651– 1652 Rikenellaceae (family), 967 Rilpivirine, for human immunodeficiency virus (HIV), 1870, 1874 Rilpivirine resistance, 1897–1898 Rimantadine, 1471, 1886–1887, 1916 Rimantadine resistance, 1903, 1917, 1921 Ring-stage survival assay, 2565 Ringworm, 2135–2136 Rio Mamoré virus, 1661 RipSeq, 580 RipSeq Mixed, 39 Risk management, laboratory safety and, 170 Risk-based classification of microorganisms, 171 Ritonavir for hepatitis C virus, 1879–1880 for human immunodeficiency virus (HIV), 1871, 1876 resistance, 1898 Ritter’s disease, 360 Rituximab Epstein-Barr virus, 1746 human herpesvirus 8 (HHV-8), 1763 multicentric Castleman’s disease, 1763 primary effusion lymphoma (PEL), 1763 River blindness, 2466 R-Mix adenoviruses, 1426 immunofluorescence detection of respiratory pathogens, 1426 influenza viruses, 1426, 1479 parainfluenzaviruses, 1426, 1491–1492 respiratory syncytial virus, 1426, 1505 virus susceptibility profiles, 1429 R-Mix refeed and rinse medium, 1430 R-Mix Too, 1426 parainfluenza virus, 1491–1492 respiratory syncytial virus, 1505 virus susceptibility profiles, 1429 RNA probes, 54, 56 quantifying in a sample, 73 RNA polymerase, 63 mutations in, 1218, 1233, 1358–1359 rifampin action on, 1358 RNA-dependent RNA polymerase hepatitis C virus, 1599, 1901–1903 rhinoviruses, 1551–1552 RNAlater, 1411 RNAmmer, 233 RNase, 1411 RNase inhibitors, 1411 RND (resistance-nodulation-cell division) type efflux pumps, 1218–1220, 1234 ROB-1, 1320, 1331 Robert Koch Institute, 150 Robinsoniella, 921 Robinsoniella peoriensis, 924–925, 930 Rochalimaea, 873 Roche 454 platform, 70 Roche Elecsys HSV-1 IgG and HSV-2 IgG assays, 1693 Rocio virus, 1645 Rocky Mountain spotted fever (RMSF), 1124–1126, 1129–1130, 2507, 2513, 2521 Romana’s sign, 2362 Roridin A, 2192 Roridin E, 2190 Rosai-Dorfman disease, 1756

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 135

Rosco Neo-Sensitabs, 1300 Roseburia, 921 Roseburia intestinalis, 922 Roseola, 1755–1756, 1760–1761 Roseolovirus (genus), 1398, 1754, 1761 Roseomonas, 829–830 Roseomonas cervicalis, 827, 830 Roseomonas fauriae, 830 Roseomonas genomospecies 4, 5, 827, 830 Roseomonas gilardii, 830 Roseomonas gilardii subsp. gilardii, 827, 830 Roseomonas gilardii subsp. rosea, 827, 830 Roseomonas mucosa, 827, 830 Ross River virus, 1645, 1651, 1655 Rotarix, 1622 RotaTeq, 1622 Rotavirus (genus), 1399, 1617 Rotaviruses antigen detection, 1623–1624 antigenic and genetic typing systems, 1628 cell culture, 1627 clinical significance, 1620–1622 description of agents, 1618 detection and identification methods, 1435 electron microscopy, 1619, 1623 epidemiology and transmission, 1620 evaluation, interpretation, and reporting of results, 1628–1629 molecular detection assays, 1625–1627 PCR, 1626 serologic tests, 1628 specimen collection and handling, 1406, 1415 taxonomy, 1617–1618 vaccine, 1622 Rothia, 354, 356 antimicrobial susceptibility testing, 1328 clinical significance, 479 description of genus, 477 epidemiology and transmission, 478 identification, 438, 494–495 isolation procedures, 480 taxonomy, 474–475 Rothia aeria, 477, 494 Rothia dentocariosa, 476–478, 484, 486, 489, 494–495 Rothia mucilaginosa, 354–357, 361–362, 364, 366–367, 371, 422, 427, 428, 429, 477, 494 Rotor-Gene, 74 RPMI 1640 medium, 1430 rpoB gene, 429 anaerobic Gram-negative rods, 983 Brucella, 1325 Corynebacteria, 484 Helicobacter, 1023 Legionella pneumophila, 898 Mycobacterium, 580, 603, 1358–1359 RPS adenovirus detector, 1774 rpsL gene, 1358, 1360 rRNA sequence analysis, 260–261 rrs gene aminoglycoside resistance, 1358, 1360 Anaplasma, 1135–1136, 1143 capreomycin resistance, 1360 Ehrlichia, 1135–1136, 1140 Orientia tsutsugamushi, 1122 RsiP gene, 1324 RSV, see Respiratory syncytial virus RSV Duet stain, 1427 RSV Respi-Strip, 1504 RSV+hMPV molecular assay, 1506 RT-helicase-dependent amplification (HDA), influenza viruses, 1476

n cxxxv

RT-nicking enzyme amplification reaction (RT-NEAR), influenza viruses, 1476 RT-PCR, 57–58 arboviruses, 1648–1652, 1652 arenaviruses, 1676–1677 coronaviruses, 1567, 1569–1577 enteroviruses, 1542–1543 filoviruses, 1676–1677 hantaviruses, 1663, 1665 hepatitis C virus, 1603–1607 hepatitis E virus, 1588, 1591–1592 human herpesvirus 6 (HHV-6), 1759 human immunodeficiency virus, 1442– 1443, 1447, 1450–1451 human metapneumovirus, 1510–1512 influenza viruses, 1476–1478, 1480–1482 Marburg virus, 1673 measles virus, 1522, 1525 mumps virus, 1494 rabies virus, 1637, 1640 respiratory syncytial virus, 1502, 1506– 1508, 1510–1512 rhinoviruses, 1552, 1554–1555, 1557–1559 rubella virus, 1527–1528, 1530 RT-SDA, 65 Rubella, 1526–1527 congenital rubella syndrome (CRS), 1526–1530 epidemiology and transmission, 1526 laboratory tests suggested for, 125 postnatal, 1526–1530 timing of biological markers of infection, 1527 TORCH (toxoplasmosis, other, rubella, cytomegalovirus, and herpes simplex virus) panels, 1530 Rubella virus, 1525–1539 clinical significance, 1526–1527 description of agent, 1525–1526 detection and identification methods, 1435 direct examination, 1527–1528 ELISA, 1528–1529 epidemiology and transmission, 1526 evaluation, interpretation, and reporting of results, 1530 hemagglutination inhibition (HI) test, 1529 immunocolorimetric assay (ICA), 1528– 1530 immunofluorescence assay (IFA), 1528– 1530 isolation and identification, 1527–1528 latex agglutination tests, 1529 nucleic acid detection, 1527 plaque reduction neutralization (PRN), 1529–1530 prenatal screening, 1530 sequencing, 1528 serologic tests, 1528–1530 specimen collection and handling, 1406– 1407, 1413–1414, 1527 taxonomy, 1525 vaccine, 1526 Rubivirus (genus), 1399, 1525 Rubulavirus (genus), 1398 Ruminococcus gauvreauii, 910 Ruminococcus lactaris, 1381 RUO (research use only) products, 82 RVAs, 1915 Saaremaa virus, 1660–1661, 1664 SabHI medium, 1951–1952 Sabia virus, 1669, 1671, 1674 Sabouraud brain-heart infusion (SABHI), 1961–1962

cxxxvi

n

SUBJECT INDEX

Sabouraud dextrose agar (SDA), 1951–1952, 1962 Sabouraud glucose agar (SGA), 2138–2139 Saccharomonospora chemotaxonomic and lysosome growth characteristics, 509 description of genus, 509, 1988 identification, 438 morphologic characteristics, 508 taxonomy, 505 Saccharomyces, 1937, 2001 clinical significance, 1994 description, 1988 microscopic appearance on morphology agar, 1998 taxonomy, 1985 Saccharomyces boulardii, 944, 1985 Saccharomyces cerevisiae, 1991, 2000 antifungal resistance, 2238, 2240 antifungal susceptibilities, 2005 ascospores, 2000 clinical significance, 1994 cultural and biochemical characteristics, 1989 description, 1988–1990 genome sequence, 241 media, 1959 taxonomy, 1985 Saccharomycetales (order), 1937, 1938, 1984 Saccharomycetes (class), 1937–1938 Saccharopolyspora chemotaxonomic and lysosome growth characteristics, 509 description of genus, 509 identification, 438, 523 morphologic characteristics, 508 taxonomy, 505 Safety equipment, 172–174; see also Laboratory biosafety biosafety cabinets, 172, 173 centrifuges, 174 chemical fume protection, 172–173 installation, 173 medical waste, 174 sharps protection, 174 splashguards, 172 Safety preparedness, 174–175 Sagenomella chlamydospora, 2064, 2073 Sagenomella sclerotialis, 2073 Saksenaea, 2088, 2091, 2094 Saksenaea erythrospora, 2088, 2094 Saksenaea oblongispora, 2088, 2094 Saksenaea vasiformis, 1962, 2088, 2094, 2096, 2099 Saksenaeaceae (family), 2088 Saline, 1423 Salinivibrio, 762 Salinococcus, 354, 356–357, 361 Salmonella, 699–705 antibiotic resistance, 1224 antimicrobial susceptibilities, 704–705, 1172–1173, 1179–1180, 1183, 1186, 1195 antimicrobial susceptibility testing, 1270 β-lactamases, 1299 biochemical reactions, 687 clinical significance, 701–702 commercial sources of chromogenic agar media for, 326 description of genus, 700–701 direct examination, 702 epidemiology and transmission, 701 evaluation, interpretation, and reporting of results, 705

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 136

identification, 702–703 H antigen determination, 703–704 MALDI-TOF MS, 703 O antigen determination, 703 phenotypic, 702–703 serogrouping and serotyping, 703–704 Vi antigen determination, 704 isolation procedures, 702 mannitol selenite broth for, 339 mannitol-lysine-crystal violet-brilliant green agar for, 339 molecular serotyping, 145 selenite cystine broth for, 344 serologic tests, 704 serotypes, 701 specimen collection, transport, and storage guidelines, 301–302 subtyping, 138, 141 taxonomy, 699–700 tetrathionate broth, Hajna, 345 typing systems, 704 Salmonella bongori, 700–701 Salmonella enterica, 700–701 Salmonella enterica serotype Choleraesuis, 702, 704 Salmonella enterica serotype Dublin, 702, 704 Salmonella enterica serotype Enteritidis, 701, 703–704 Salmonella enterica serotype Gallinarum, 701 Salmonella enterica serotype Paratyphi, 344, 1254 Salmonella enterica serotype Paratyphi A, 700, 702–703, 705 Salmonella enterica serotype Paratyphi B, 702, 704–705 Salmonella enterica serotype Paratyphi C, 702, 704–705 Salmonella enterica serotype Senftenberg, 1227 Salmonella enterica serotype Typhi, 344, 700– 705, 1173, 1193, 1254 Salmonella enterica serotype Typhimurium, 703, 1216, 1229 Salmonella enterica subsp. arizonae, 700–701 Salmonella enterica subsp. diarizonae, 700–701 Salmonella enterica subsp. enterica, 700–701 Salmonella enterica subsp. houtenae, 700–701 Salmonella enterica subsp. indica, 700–701 Salmonella enterica subsp. salamae, 700–701, 703 Salmonella-shigella agar, 343 Salmonellosis, laboratory tests suggested for, 125 Salpingitis Chlamydia trachomatis, 1108 etiologies, usual, 290 Mycoplasma, 1092 Neisseria gonorrhoeae, 636 Salt meat broth, 343 Salt tolerance medium, 343 Sandflies, 2505–2506 Sandfly fever virus, 1645, 2507 Sapovirus (genus), 1399, 1617 Sapoviruses cell culture, 1627 clinical significance, 1620 description of agents, 1619 electron microscopy, 1619, 1623 epidemiology and transmission, 1620–1621 molecular detection assays, 1625–1627 PCR, 1626 taxonomy, 1617–1618 Sappinia, 2327, 2387 Sappinia diploidea, 2387

Sappinia pedata, 2387 Saprochaete capitata, 1984 Saprophytic fungi, 1951 Saquinavir, for human immunodeficiency virus (HIV), 1871, 1876 Saquinavir resistance, 1897–1898 Sarcinomyces phaeomuriformis, 2154 Sarcocystidae (family), 2373, 2425 Sarcocystis, 2425–2431 antigen detection, 2430 clinical significance, 2429 collection, transport, and storage of specimens, 2429 culture, 2430 description of agents, 2426–2427 detection, 2329, 2332 direct examination, 2429–2430 epidemiology, transmission, and prevention, 2428 evaluation, interpretation, and reporting of results, 2431 life cycles, 2427 microscopy, 2426–2427, 2429–2430 muscular infection, 2427–2431 nucleic acid detection, 2430 serologic tests, 2430 stains for detection, 2312 taxonomy, 2425 treatment, 2431 Sarcocystis hominis, 2425, 2427–2431 Sarcocystis suihominis, 2425, 2427–2431 Sarcodina (subphylum), 2387 Sarcophaga, 2519 Sarcophagidae (family), 2513 Sarcoptes scabiei, 2516–2517 Sarcoptic mange, 2516 Sarocladium, 2064, 2071, 2076 Sarocladium bacillisporum, 2064, 2071 Sarocladium kiliense, 1967, 2064, 2071, 2177 Sarocladium strictum, 2064, 2071 SARS (severe acute respiratory syndrome), 120, 147, 1565–1578 SARS-CoV, 1565–1578 antigen detection, 1570 biosafety, 1570, 1577 clinical significance, 1569 collection, transport, and storage of specimens, 1570 description of agent, 1565–1566 direct detection, 1570–1577 discovery, 1566 epidemiology and transmission, 1567, 1569 evaluation, interpretation, and reporting of results, 1578 isolation procedures, 1577 nucleic acid detection, 1571 origin, 1565 phylogenetic relationships, 1566 serologic tests, 1577 structure, 1565–1566 taxonomy, 1565–1566 SAS Adeno test, 1624 SAS FluAlert Influenza A test, 1474 SAS FluAlert Influenza A&B test, 1474 SAS FluAlert Influenza B test, 1474 SAS HMPV test, 1510 SAS rapid adenovirus test, 1774 SAS Rota test, 1624 SAS RSV, 1504 SaSelect medium, 343 Satratoxins, 2190 Scabies, 2516–2517, 2534 Scalars, arthropods as

SUBJECT INDEX cockroaches, 2513, 2515 muscoid flies, 2513 Scalded skin syndrome, staphylococcal, 360, 365–366 Scardovia, 920–921, 930 Scardovia inopinata, 925 Scardovia wiggsiae, 925–926 Scarlet fever, 385 Scedosporiosis, 2167 Scedosporiosis, specimens for, 1947 Scedosporium, 1937, 1939, 1940, 1950, 2057, 2153, 2155, 2159, 2162–2168, 2173– 2174, 2176, 2178, 2180–2181 antifungal resistance, 2243, 2245 antifungal susceptibility testing, 2271– 2272 microscopy, 1967, 1969 Scedosporium apiospermum, 2118, 2155, 2159, 2163, 2167, 2173, 2177, 2181 antifungal susceptibility testing, 2257, 2262, 2268–2269, 2271–2272 microscopy, 1965, 1975 Scedosporium aurantiacum, 2155, 2159, 2167, 2257 Scedosporium boydii, 1938, 2155–2156, 2159, 2167, 2173–2174, 2176–2177, 2180– 2182 Scedosporium prolificans, 2073, 2155, 2159, 2163, 2166–2167, 2271–2272 Scedosporium selective medium, 1952 SCGYEM medium, 2315 Schaedler agar, 343 Schaedler CNA agar with vitamin K1 and sheep blood, 343 Schaudinn’s fixative/solution, 2303–2304, 2311–2312 Schistosoma, 2483 antiparasitic agent resistance, 2555–2556 commercial kits for immunodetection of serum antibodies, 2296 detection, 2323, 2329 treatment, 2531, 2533 Schistosoma bovis, 2480 Schistosoma curassoni, 2480 Schistosoma guineensis, 2480 Schistosoma haematobium, 2479–2480, 2483– 2486 detection, 2320, 2326, 2328, 2330 eggs, 2449 specimens, 2305 Schistosoma intercalatum, 2320, 2480, 2483– 2485 Schistosoma japonicum, 2479–2480, 2483– 2486 detection, 2320, 2329, 2332 eggs, 2449 Schistosoma malayensis, 2480 Schistosoma mansoni, 2479–2480, 2483, 2485–2486 antiparasitic agent resistance, 2555–2556 detection, 2320, 2329, 2332 eggs, 2449 Schistosoma margrebowiei, 2480 Schistosoma mattheei, 2480 Schistosoma mekongi, 2320, 2479–2480, 2483, 2485 Schistosoma sinensium, 2480 Schistosoma spindale, 2484 Schistosomatidae (family), 2290, 2479 Schistosomatoidea (superfamily), 2290 Schistosomes, 2479–2486 anthelminthic susceptibility and treatment, 2486 avian, 2486

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 137

clinical significance, 2484–2485 collection, transport, and storage, 2486 dermatitis (swimmer’s itch), 2480, 2486 detection, 2486 epidemiology and transmission, 2484 hatching of eggs, 2323 serology, 2486 taxonomy, 2479–2480, 2484 typing, 2486 Schistosomiasis acute toxemic (Katayama fever), 2484 antiparasitic agent resistance mechanisms, 2555–2556 antiparasitic agent susceptibility testing methods, 2564, 2567 commercial kits for immunodetection of serum antibodies, 2296 genitourinary, 2480, 2484–2485 hepatosplenic, 2479, 2484 intestinal, 2480 rectal, 2480 treatment, 2533 Schizophyllum, 1937, 2063, 2069–2071, 2075 Schizophyllum commune, 1938, 2063, 2070– 2071, 2075, 2076, 2077 Schizopyrenida (order), 2287, 2387 Schizosaccharomyces, 2015 Schizosaccharomyces pombe, 2245 Schizotrypanum (subgenus), 2357 Schleifer-Krämer agar, 343 Schneider’s Drosophila medium with 30% fetal calf serum, 2315 Schüffner dots, 2335, 2343, 2345 Sclerosing cholangitis, Cryptosporidium, 2438 Sclerotic body, 1942, 1967, 1975 Sclerotium, 1942 Scolecobasidium gallopavum, 2268–2269 Scolex, 2324, 2332, 2471–2475, 2477 Scolopendra, 2520 Scopulariopsis, 1940, 2069, 2071, 2075, 2077, 2159 cycloheximide inhibition, 1951, 1955 key phenotypic features, 2065 Scopulariopsis acremonium, 2065, 2075 Scopulariopsis brevicaulis, 2065, 2075, 2077 Scopulariopsis brumptii, 2075, 2167 Scopulariopsis candida, 2065, 2075 Scorpion probes, 61, 62 Scorpiones (subclass), 2522 Scorpions, 2519–2520, 2522 Scrofula, 544 Scrub typhus, 92, 1124–1125, 1129–1130, 2507, 2511 Scutigera coleoptrata, 2520 Scytalidium, 2073, 2076, 2146 Scytalidium cuboideum, 2073, 2076 Scytalidium dimidiatum, 1939, 2057, 2153 Scytalidium hyalinum, 2153 SD Bioline, 1069 SDD (susceptible dose-dependent), 2257 Seadornavirus (genus), 1399 Sealpox, 1828, 1830 Secernentea (class), 2289, 2448, 2454, 2461, 2465, 2468, 2493, 2495–2499 Secnidazole Dientamoeba fragilis, 2413 Giardia duodenalis, 2412 Sedimentation for cestodes, 2473–2476 for schistosomes, 2486 ova and parasite (O&P) examination, 2304–2305 Sedoreovirinae (subfamily), 1399 Seegene RV12 ACE detection assay, 1478

n cxxxvii

Seegene RV15 ACE detection assay, 1478 Seeplex HPV 6, 1792 Seeplex HPV-4 ACE, 1790 Seeplex HPV-18, 1792 Seeplex respiratory virus detection assay, 1511 Seeplex RV15, 1575 Seeplex RV15 OneStep ACE detection, 1555 Seeplex STD 6 ACE detection system, 2415 Segniliparus clinical significance, 519 description of genus, 509, 511 G+C content, 536 identification, 438 microscopy, 521 morphologic characteristics, 508 taxonomy, 504–505 Segniliparus rotundus, 512, 514, 519 Segniliparus rugosus, 510, 512, 514, 519, 527 Seizures carbapenems, 1177 herpes B virus, 1697 human herpesvirus 6 (HHV-6), 1755– 1756 human herpesvirus 7 (HHV-7), 1761 influenza virus, 1471 penicillins, 1173 phaeohyphomycoses, 2161 quinolones, 1180 Rickettsia, 1124 Taenia solium, 2476 Selenite broth (selenite-F broth), 343 Selenite broth base, mannitol, 344 Selenite cystine broth, 344 Selenomonas, 974 characteristics of genus, 970–971 clinical significance, 974 identification, 977, 981–982 taxonomy, 969 Selenomonas artemidis, 969, 982 Selenomonas dianae, 969, 982 Selenomonas flueggei, 969, 982 Selenomonas infelix, 969, 982 Selenomonas noxia, 969, 974, 982 Selenomonas sputigena, 969, 974, 982 Sellers stain, 1638–1639 Semiapochromates, 8 Semliki Forest virus, 1645 Sensititre Streptococcus, 1320 Streptococcus pneumoniae, 1319 Sensititre ANO2B, 1345 Sensititre ARIS, 34, 1277 Sensititre ID plates, 34, 1277 Sensititre MycoTB panel, 586, 1277, 1367 Sensititre Vizion system, 1275 Sensititre Windows software (SWIN), 34, 1275, 1277 Sensititre Yeast YO10 system, 2045 Sensititre YeastOne, 2264–2265, 2271–2272 SensiTrop II HIV coreceptor tropism assay, 1448 SENTRY antimicrobial surveillance program echinocandin resistance, 2244 Streptococcus pneumoniae, 1315–1316 Seoul virus, 1660–1662, 1664 Sepedonium, 2118 Sepsis Bacillus pumilus, 443 Campylobacter, 1000 Citrobacter, 720 Clostridium, 948 Corynebacterium amycolatum, 479

cxxxviii n

SUBJECT INDEX

Sepsis (continued) Fusobacterium, 973 herpes simplex virus (HSV), 1689 Neisseria meningitidis, 637 Pediococcus, 424 Plesiomonas shigelloides, 721 Pseudomonas aeruginosa, 775 Sneathia, 974 Sphingobacterium, 825 Streptococcus bovis group, 387 Streptococcus mitis group, 386 surrogate markers for, 22–23 SepsiTest, 1381 Septata intestinalis, 2209–2210 Septate, 1935, 1938, 1939, 1942 Septic abortion Gram-positive anaerobic cocci (GPAC), 910 Leptotrichia, 974 Septic arthritis Abiotrophia and Granulicatella, 424 Burkholderia, 794 Finegoldia magna, 911 Fusarium, 2058 Gemella, 424 Haemophilus haemolyticus, 670 Mycobacterium malmoense, 542 Mycobacterium xenopi, 543 Neisseria meningitidis, 637 Pantoea, 719 Septic shock Gemella, 424 meningococcal, 637 Plesiomonas shigelloides, 721 Septicemia Achromobacter group B, 824 Aeromonas, 754 Bacillus cereus, 443, 1326 Bacillus clausii, 442 Bacillus licheniformis, 442 Bacillus subtilis, 442 Bergeyella zoohelcum, 827 Burkholderia, 794 Capnocytophaga, 654 Citrobacter, 720 Dolosigranulum pigrum, 424 Edwardsiella, 721 Elizabethkingia meningoseptica, 828 Fusobacterium, 973 Lactococcus, 424 Leptospira, 1030 Listeria monocytogenes, 463 Methylobacterium, 830 Micrococcaceae, 361 Moraxella, 814 Neisseria lactamica, 645 Neisseria weaveri, 646 Parvimonas micra, 911 Pasteurella, 655 Rhodotorula, 1994 Rothia mucilaginosa, 361 Serratia, 720 Shewanella, 825 Vibrio vulnificus, 765 Yersinia enterocolitica, 742 Septicemic plague, 741–742 Septi-Chek, 20, 1948 SeptiFast, 23, 1979 Septum, 1935, 1939, 1942 SeqHepB database, 1921 SeqNet, 150 Sequence analysis of protein-encoding genes, 261 Sequence database, Mycobacterium, 580–581

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 138

Sequence-independent single primer amplification (SISPA), 242 Sequencing, see also DNA sequence analysis for identification adenovirus, 1776 antiviral susceptibility testing, 1916–1918 Epstein-Barr virus, 1743 hantaviruses, 1664 hepatitis C virus, 1605, 1607 influenza viruses, 1480–1482 respiratory syncytial virus, 1505 rhinoviruses, 1557 rubella virus, 1528 Sequenom MassArray, 145 Serine β-lactamases, 1223 Serine carbapenemases, 1225 Serion ELISA antigen Candida, 1996 Serion ELISA classic Candida albicans, 1996 SeroCT, 1116 Serologic tests, see specific organisms Serotyping, molecular, 145 Serozyme assays, for gastroenteritis viruses, 1624 Serpulina, 1055 Serratia antimicrobial susceptibilities, 1174–1175, 1177, 1180, 1181, 1193, 1196 β-lactamases, 1299 biochemical characteristics of genus members, 724 epidemiology, transmission, and clinical significance, 720 identification, 725 taxonomy, 715 Serratia entomophila, 717, 724 Serratia ficaria, 717, 724 Serratia fonticola, 717, 724 Serratia grimesii, 717 Serratia liquefaciens, 717, 720, 724–725 Serratia marcescens, 720 antibiotic resistance, 1217, 1224–1226 antimicrobial susceptibilities, 727–730, 1174, 1191, 1195 β-lactamases, 1299 product contamination by, 126–127 Serratia marcescens subsp. marcescens, 717, 724 Serratia marcescens subsp. sakuensis, 715, 717, 724 Serratia nematodiphila, 717 Serratia odorifera, 715, 717, 724 Serratia plymuthica, 715, 717, 720, 724 Serratia proteamaculans, 717 Serratia rubidaea, 715, 717, 720, 724 Serratia ureilytica, 715, 717 Serum, antibacterial agent concentrations in, 1197–1199 Serum agglutination test, Brucella, 867–868 Serum glucose agar (serum dextrose agar), 344 Serum neutralization, see Neutralization assay Serum sickness, cephalosporins and, 1175 Serum tellurite agar, 344 Sessile, 1942 Setosphaeria rostrata, 2155 SET-RPLA kit, 366 Severe acute respiratory syndrome (SARS), 120, 147, 1565–1578 Severe asthma with fungal sensitization (SAFS), 2044 Sexual reproduction, in fungi, 1936–1938 Sexual transmission adenovirus, 1772

herpes simplex virus (HSV), 1688–1689 human herpesvirus 8 (HHV-8), 1762 human papillomavirus (HPV), 1783–1785 molluscum contagiosum virus, 1828–1829, 1831 pubic louse, 2511 Trichomonas vaginalis, 2413–2415 Shannon’s index, 134 Sharps protection, 174 Sheep tick, 2516 Shell vial centrifugation culture (SVCC), for adenovirus, 1775–1776 Shepard’s differential agar, 325 Sherlock system, 34–35 coryneform Gram-positive rods, 483 Staphylococcus, 364 Shewanella, 615, 825 Shewanella algae, 624–625, 823, 825 Shewanella putrefaciens, 624–625, 823, 825 Shiga toxin-producing Escherichia coli (STEC) clonality, 148 specimen collection, transport, and handling, 302–303 subtyping, 138 Shigella, 697–699 antibiotic resistance, 1224 antimicrobial susceptibilities, 699, 1172, 1180, 1183, 1186, 1192, 1195 antimicrobial susceptibility testing, 1270 β-lactamases, 1299 clinical significance, 698 description of genus, 697 direct examination, 698 epidemiology and transmission, 697–698 Escherichia coli compared to, 685, 697 evaluation, interpretation, and reporting of results, 699 identification, 698–699 phenotypic, 698–699 serotyping, 699 isolation procedures, 698 serological tests, 699 specimen collection, transport, and storage guidelines, 301–302 taxonomy, 697 typing systems, 699 Shigella boydii, 685, 687, 693, 697–699 Shigella dysenteriae, 687, 697–699 Shigella flexneri, 687, 697–699 Shigella sonnei, 687, 697–699 Shigellosis, 125, 2513 Shimoni bat virus, 1633–1634 Shimwellia blattae, 685 Shingles, see Varicella-zoster virus Short gut syndrome, Leuconostoc and, 424 Shuttleworthia, 921 Shuttleworthia satelles, 924, 930 SHV β-lactamases, 1223–1225, 1299 Siccibacter turicensis, 719 Sick building syndrome, 2192 Siemens Advia Centaur HIV 1/O/2, 1444 σ-Swab, 48 Sigma VCM medium, 1410 Sigma Virocult medium, 1410 Sigmoidoscopy specimens, 2324–2325 Signal amplification techniques, 55–57 bDNA assays, 55–56 Cleavase-Invader technology, 56–57, 58 hybrid capture assays, 56, 57 Signaling-lymphocytic activation molecule (SLAM), mumps virus, 1494 Silver ions, for disinfection, 195 Silver staining, for Treponema pallidum, 1063, 1073

SUBJECT INDEX Simeprevir, for hepatitis C virus, 1601–1602, 1879 Simeprevir resistance, 1902 Simian immunodeficiency virus (SIV), 1436 Simian T-cell lymphotropic viruses (STLVs), 1458–1459 Simian virus 40 (SV40), 1803–1806 Simmons’ citrate agar, modified, 327 Simmons’ citrate agar (citrate agar), 344 Simonsiella antimicrobial susceptibilities, 662 clinical significance, 655 direct examination, 656 epidemiology and transmission, 654 identification, 660 isolation procedures, 656, 658 taxonomy and description of, 653 Simonsiella muelleri, 653, 655–656, 658, 660, 662 Simple microscope, 7 Simplexa CMV, 1726 Simplexa Flu A/B & RSV, 1477, 1506 Simplexa Flu A/B & RSV Direct, 1506 Simplexvirus (genus), 1398 Simpson’s index of diversity (DI), 134 SimulFluor reagents, 1489 SimulFluor Respiratory Screen RSV/flu A, 1503 SimulFluor Respiratory Screen RSV/Para 3, 1503 Simulium, 2507 Sin Nombre virus, 1661, 1665 Sindbis virus, 1645, 1652 Single nucleotide polymorphism (SNP) antiviral susceptibility testing, 1916, 1918 DNA microarray analysis, 145 k-mer analysis, 144 mass spectrometry analysis, 145 Pseudomonas, 780 varicella-zoster virus, 1711 whole-genome SNP, typing, 141–143 Single-strand conformation polymorphism (SSCP) analysis, 67 Single-use devices, reuse of, 205–206 Single-vial collection systems, for stool specimen preservation, 2303–2304 Single-well broth dilution method for clindamycin resistance detection Staphylococcus, 1267, 1296 Streptococcus, 1267, 1298 for mupirocin resistance detection in Staphylococcus, 1297 Sinomonas, 354, 361 Sinus specimen collection, transport, and handling, 271, 300 fungi, 1946–1947 parasitology, 2329, 2332 Sinusitis Chlamydia pneumoniae, 1108 Coniochaeta hoffmannii, 2076 Finegoldia magna, 911 Fusarium, 2058 Fusobacterium, 973 Haemophilus influenzae, 669 Haemophilus parainfluenzae, 670 hyaline fungi, 2075–2076 influenza virus, 1471 microsporidia, 2210, 2213 Moraxella, 814 mucormycosis, 2089 Parvimonas micra, 911 phaeohyphomycoses, 2161, 2163–2164 rhinoviruses, 1553

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 139

Schizophyllum commune, 2075 Scopulariopsis, 2075 Staphylococcus, 360 Siphonaptera (order), 2507, 2509–2510, 2522 SIR Mycoplasma, 1099 SIRE supplement, 1365–1366 SIV (simian immunodeficiency virus), 1436 Six Sigma, 47 16S rRNA gene sequence, 39, 75–76 Achromobacter, 843 Aeromonas, 756 Alcaligenes, Kerstersia, Advenella, and Paenalcaligenes, 843 anaerobic Gram-negative rods, 969, 974– 975, 979, 982–983 Anaplasma, 1135–1136, 1143 Arcobacter, 1005–1006 Bacillus, 454 bacteria classification and, 257, 260–261 Bartonella, 877 Bordetella, 843 Borrelia, 1037 Brachyspira, 1064 Brucella, 865 Burkholderia, 796, 799 Campylobacter, 1005–1006 catalase-negative, Gram-positive cocci, 428–429 Chlamydiaceae, 1106, 1112, 1114 Clostridium, 940, 948, 954–955 Corynebacterium, 484 Cupriavidus, 800 Ehrlichia, 1135–1136, 1140 Enterobacteriaceae (family), 714 Enterococcus, 411 Escherichia coli, 694 Eubacterium, 922 Francisella, 858 HACEK group, 658, 660–661 Haemophilus, 672 Helicobacter, 1013, 1015, 1019, 1023 identification of Gram-positive cocci, 352 Legionella pneumophila, 892, 898 microbiome analysis and, 226, 229, 231 Mycoplasma pneumoniae, 1094 Nocardia, 524–525 non-spore-forming, anaerobic, Grampositive rods, 929–930 nontuberculous mycobacteria, rapidly growing, 602 nontuberculous mycobacteria (NTM), slowly growing, 579–580 Orientia tsutsugamushi, 1122 Pandoraea, 801 pathogen discovery and, 240 Pseudomonas, 776 Ralstonia, 791, 800 sequencing kits, 76 Staphylococcus, 365 Streptococcus, 393 Tropheryma whipplei, 1159, 1163 Varibaculum cambriense, 920 Vibrionaceae, 768 16S rRNA methylases, 1383 Skim milk, 320–321 Skin and soft tissue infection (SSTI) anaerobic Gram-negative rods, 972 clostridial histotoxic, 944–946 Dialister, 974 Finegoldia magna, 910–911, 911 Gram-negative curved bacilli, 997 Gram-positive anaerobic cocci (GPAC), 910–911

n cxxxix

Mycobacterium marinum, 1370 non-spore-forming, anaerobic, Grampositive rods, 923 Porphyromonas, 971 Prevotella, 973 Shewanella, 825 Staphylococcus, 357 Skin disinfection, 17, 187–188 Skin infection/lesions, see also Cutaneous infection/lesions Rash; Skin and soft tissue infection arthropods and, 2515, 2518–2521 Bacillus megaterium, 443 Bartonella, 876 Blastomyces dermatitidis, 2114 centipede bites, 2520 dermestid beetles and, 2521 Fusarium, 2058 Gnathostoma, 2497–2498 herpes simplex virus (HSV), 1688–1689 Lacazia loboi, 2197–2198 Lagenidium, 2198, 2204 lymphatic filariasis, 2462–2463 Malassezia, 1994 Metarhizium anisopliae, 2076 Methylobacterium, 830 microsporidia, 2210, 2213 Mycobacterium simiae, 543 Mycobacterium ulcerans, 543 Mycobacterium xenopi, 543 Onchocerca volvulus, 2466 Parvimonas micra, 911 phaeohyphomycoses, 2162, 2164 poxviruses, 1830–1831 Pythium insidiosum, 2198, 2201 Rhinosporidium seeberi, 2198, 2205 Streptococcus pyogenes, 385 Talaromyces marneffei, 2046 Treponema, 1061 trichodysplasia spinosa, 1805–1807 Trichosporon, 1994 Tropheryma whipplei, 1161 urticating caterpillar hairs, 2518–2519 varicella-zoster virus, 1704–1705 yatapoxviruses, 1831 Skin microbiome, 231 Skin specimen/biopsy fungi, 1945, 1947, 1949 Orientia, 1127 parasitology, 2294, 2300, 2329, 2332–2333 Rickettsia, 1126 viruses, 1415 Skinner Tank virus, 1669, 1672 Skirrow brucella medium, 344 Slackia, 920–921 Slackia exigua, 925, 930 Slackia heliotrinireducens, 910 SLAM (signaling-lymphocytic activation molecule), mumps virus, 1494 Slapped-cheek disease, 1819, 1822 Sleeping sickness, 2357, 2366–2368, 2506, 2507, 2555 Slidex Staph Plus, 363 Slipped-strand mispairing (SSM), 140 Small bowel bacterial overgrowth syndrome, 303 Small intestinal microbiome, 230 Smallpox, 220–222, 1829–1832 Smart CMV, 1726 SmartCycler, 74 SmartGene IDNS, 580, 2003 Smudge cells, 1423 Smuts, 2075 Sneathia, 497 characteristics of genus, 970–971 clinical significance, 974 identification, 980–981 taxonomy, 968

cxl

n

SUBJECT INDEX

Sneathia amnii, 968, 974–975, 980–981 Sneathia sanguinegens, 968, 974–975, 979, 981 Sneezing, Linguatula serrata and, 2516 Société Française de Microbiologie (CASFM), 1268–1269 Sodium acetate-acetic acid-formalin (SAF), 2302–2303, 2311–2312 Sodium hippurate broth (hippurate broth), 344 Sodium hydroxide, 559, 1956 Sodium polyanethol sulfonate (SPS), 17, 497 Sodium stibogluconate, 2362, 2542, 2554, 2564 Sodium stibogluconate resistance, 2554 Sofia Influenza A+B FIA, 1474 Sofia RSV, 1504 Sofosbuvir, for hepatitis C virus, 1601–1602, 1879–1881 Sofosbuvir resistance, 1902 Soft tissue infection, see also Skin and soft tissue infection Actinobacillus, 654 Actinomyces massiliense, 924 Bacteroides, 971 Campylobacter, 1001 Capnocytophaga, 654 Coccidioides, 2114 Fusobacterium, 973 Histoplasma capsulatum, 2114 Mycobacterium, 596, 598 non-spore-forming, anaerobic, Grampositive rods, 923 Pantoea, 719 Pasteurella, 655 Proteus, 720 Stenotrophomonas maltophilia, 794 Soil extract agar, 1962 Solenopsis invicta, 2518 Solid organ transplant patients Acanthamoeba, 2389, 2392 adenoviruses, 1771 cytomegalovirus and, 1719 Epstein-Barr virus, 1740, 1746–1747 Fusarium, 2065, 2067 hepatitis E virus, 1590 Histoplasma capsulatum, 2114 human herpesvirus 6 (HHV-6), 1756 mucormycosis, 2088–2089 respiratory syncytial virus and, 1501 Trypanosoma cruzi, 2363 Solid-phase hybridization, 54, 55 Solithromycin, 1184 Solobacterium, 921 Solobacterium moorei, 922, 924, 930 Sorbitol-MacConkey agar with 5-bromo-4chloro-3-indolyl-β-D-glucuronide (BCIG), 344 Sordariales (order), 1938, 2153, 2155, 2161, 2174, 2176 Sordariomycetes (class), 1937–1938 Sore mouth, 1830 Sore throat arenaviruses, 1673 coronaviruses, 1569 Corynebacterium diphtheriae, 480 Epstein-Barr virus, 1739 filoviruses, 1674 influenza virus, 1471 Sorenson pH buffers, 320 Source attribution, 146–147 Southern blotting, for polyomaviruses, 1807 Soybean casein digest broth, 17 Soybean medium with 0.1% agar, 344

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 140

“Spaghetti and meatballs” structures, 1806, 1808, 1995, 2147 Span Crystal Tp, 1069 Span Signal Spirolipin, 1070, 1072 Spanish fly, 2521 Sparfloxacin, 1179 Sparganosis, 2332, 2471, 2477 Sparganum, 2329, 2332, 2471, 2501–2503 Spasticity, HTLVs and, 1460 Special infusion broth with blood, 344 Species criteria for delineation, 256 genomic threshold for definition, 257–258 polyphasic species concept, 256–267 Species complex, 1984 Specimen collection, transport, and processing, 270–306; see also specific organisms; specific specimen types anaerobic bacteria, 281, 905 biosafety, 1409 Clostridium, 948–952 suspected C. botulinum or C. tetani infection or intoxication, 951– 952 suspected C. difficile infection, 949–951 suspected C. perfringens foodborne illness, 948 suspected enteritis necroticans (C. perfringens type C), 948–949 suspected gas gangrene or necrotizing fasciitis, 948 suspected neutropenic enterocolitis involving C. septicum, 951 collection methods aspirates, 271 by specimen type, 272–280 feces, 281 from prosthetic devices, 281 sputum, 281 swabs, 270–271 tissue, 271, 281 urine, 281 for molecular methods, 77–78 fungi, 1944–1953 general principles, 270–282 guidelines by specimen type, 272–280 health care-associated infections, 110 laboratory handling of specimens, 283–286 culture examination and interpretation, 289 documentation, 283–284 Gram stain, 285–288 initial sample handling, 285–289 labeling of specimens, 283–284 medium inoculation, 286–287, 289 molecular detection of bacteria, 289 prioritization, 285 processed at remote site, 285 rejection of specimens, 284–285 reporting results, 289 laboratory safety issues, 282–283 parasitology, 2293–2308 blood collection, 2304 body sites and possible parasites recovered, 2294 commercial kits for immunodetection in stool samples, 2295 commercial kits for immunodetection of serum antibodies, 2296 direct detection by routine methods, 2304–2308 fecal specimen collection and processing options, 2301 sample preparation and procedures by body site, 2297

stool collection, 2293–2294, 2296 stool preservation, 2300–2304 stool test ordering, 2302 selection of specimen by anatomical site, 271 transport, specimen maintenance during, 282–283 viruses, 1405–1417 collection methods and processing of specimens, 1412–1416 specimen collection, 1405, 1408–1409 specimen selection, 1405–1408 specimen storage and processing, 1410– 1412 transport conditions, 1410 transport medium, 1409–1410 volume of sample, 281–282 Specimen preservative medium, 344 Specimen processing, automated, 48–49 Specimen rejection, 284–285, 1944 Spectinomycin, 1181 Spectra MRSA, 1293 Spectral Archiving and Microbial Identification System (SARAMIS), 35 Speed-oligo Mycobacterium assay, 583 Sphaeropsidales, 1939 Spherical aberration, 5 Spherules, 1966, 1969, 1976 Sphingobacterium, 813, 823, 825 Sphingobacterium mizutaii, 624–625, 825, 826–828 Sphingobacterium multivorum, 626–627, 823, 825 Sphingobacterium spiritivorum, 626–627, 823, 825 Sphingobacterium thalpophilum, 626–627, 823, 825 Sphingomonadaceae (family), 825 Sphingomonas, 616, 626–629, 823, 825–826, 895 Sphingomonas parapaucimobilis, 825–826 Sphingomonas paucimobilis, 615, 825–826 Spiders, 2520–2522 Spin amplification shell vial assay, for cytomegalovirus, 1727–1728 Spinareovirinae (subfamily), 1399 Spinosad, 2511 Spiramycin, 1182 Cryptosporidium, 2442 Toxoplasma gondii, 2381–2382 Spirochaetaceae (family), 1037 Spirochaetales (order), 1028, 1037 Spirochaetes (class), 1028 Spirochaetes (phylum), 1028 Spirochetes, human host-associated, 1055– 1075 antimicrobial susceptibilities, 1072 Brachyspira, 1072 Treponema pallidum, 1072 clinical significance, 1059–1061 endemic treponematoses, 1061 intestinal spirochetosis, 1061 venereal syphilis, 1059–1061 collection, transport, and storage of specimens, 1061–1063 intestinal spirochetosis, 1063 oral treponemes (T. denticola), 1063 syphilis (T. pallidum), 1062–1063 dark-field microscopy, 9 description of agents, 1055–1057 direct examination, 1063–1064 Brachyspira, 1063–1064 Treponema pallidum, 1063

SUBJECT INDEX epidemiology and transmission, 1057–1059 endemic treponematoses, 1058 intestinal spirochetosis, 1059 oral treponemes, 1058–1059 venereal syphilis, 1057–1058 evaluation, interpretation, and reporting of results, 1072–1075 direct detection of Treponema pallidum, 1072–1073 serologic tests, 1073–1074 syphilis tests in HIV infection, 1075 tests for congenital syphilis, 1075 tests for neurosyphilis, 1074 identification of Brachyspira, 1064–1065 isolation procedures, 1064 Brachyspira, 1064 Treponema pallidum, 1064 serologic tests, 1065–1072 general principles, 1065–1066 nontreponemal tests, 1066, 1073–1074 treponemal tests, 1066–1072, 1074 taxonomy, 1055 treponemal tests for syphilis, 1066–1072 chemiluminescence immunoassays (CLIAs), 1071–1072 combined treponemal IgM/IgG EIAs, 1068 conventional, 1066, 1070, 1074 EIAs, 1067–1068, 1070–1071, 1074 FTA-ABS test, 1066, 1070, 1074 immunoblot assays, 1070, 1072 MHA-TP test, 1066, 1070, 1074 multiplex flow immunoassays, 1071– 1072 rapid combination nontreponemal and treponemal tests, 1070 rapid treponemal tests, 1069, 1071– 1072, 1074 TPHA, 1066, 1070 TP-PA test, 1066, 1070, 1074 treponemal IgM or IgG EIAs, 1067 typing systems, 1065 Brachyspira, 1065 Treponema pallidum, 1065 Spirometra, 2477, 2502–2503 clinical significance, 2503 description of agents, 2502 detection, 2329, 2332 direct examination by microscopy, 2503 epidemiology, transmission, and prevention, 2502–2503 serologic tests, 2503 treatment, 2503 Spirometra erinacei, 2502 Spirometra mansoni, 2502 Spirometra mansonoides, 2471, 2477, 2502 Spirometra ranarum, 2502 Spiroplasma, 1089–1090 Spiroplasmataceae (family), 1089 Spiruria (subclass), 2461, 2465, 2467–2468 Spirurida (order), 2289, 2461, 2465, 2467– 2468 Spiruroidea (superfamily), 2289 Splashguards, 172 Spleen specimen, for parasitology, 2294, 2299, 2328, 2330–2331 Splendore-Höeppli phenomenon, 1969, 2100, 2103, 2177, 2202 Splenic peliosis, 876 Splenic rupture, Plasmodium vivax, 2341 Splenomegaly Epstein-Barr virus, 1739 human herpesvirus 8 (HHV-8), 1762– 1763

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 141

Leishmania, 2359 Trypanosoma brucei, 2366 Trypanosoma cruzi, 2362 Trypanosoma lewisi, 2368 Spoligotyping, for Mycobacterium tuberculosis complex, 581, 584 Spondylodiscitis Aerococcus, 424 Aggregatibacter, 654 Bacteroides, 970 Blastoschizomyces, 1992 Lactococcus, 424 Mycobacterium xenopi, 543 Nocardia nova, 517 Sporadic endemicity, definition, 132 Sporangiole, 1942 Sporangiolum, 1942 Sporangiophore, 1937, 1942 Sporangiospores, 1937 Sporangium, 1937, 1942 Spore stain, 323 Spore test, Clostridium, 953–954 Spores, fungal, 1936–1938 Sporobolomyces antifungal susceptibility testing, 2263 clinical significance, 1994 description of agents, 1991 taxonomy, 1985 Sporobolomyces holsaticus, 1985, 1994 Sporobolomyces johnsonii, 1985 Sporobolomyces roseus, 1985, 1994 Sporobolomyces salmonicolor, 1985, 1989, 1994, 2005, 2264 Sporodochia, 2058, 2060–2061 Sporodochium, 1942 Sporolactobacillaceae, 441 Sporothrix, 1966, 2153, 2155, 2159, 2162, 2164–2167 Sporothrix brasiliensis, 2155, 2159, 2161, 2163, 2167 Sporothrix cyanescens, 2071 Sporothrix globosa, 2155, 2159, 2167 Sporothrix luriei, 2155, 2159 Sporothrix schenckii, 1935, 2155, 2159, 2161, 2166–2167 antifungal susceptibility testing, 2268– 2269 endophthalmitis, 1949 media for, 1953, 1959 microscopy, 1976 Sporotrichosis, 2153 clinical significance, 2164 epidemiology and transmission, 2161 microscopy, 2164 specimens for, 1947 Sporotrichum, 2063 Sporotrichum pruinosum, 2063, 2071 Sporozoites, Toxoplasma gondii, 2373–2374 Spumaretrovirinae (subfamily), 1399 Spumavirus (genus), 1399 Sputasol, 2331 Sputolysin, 1950 Sputum specimen Advenella, 841 Burkholderia, 795 direct wet mount, 2331 fungi, 1946, 1948, 1950 Gram stain and plating medium recommendations, 286 Kerstersia, 841 Legionella, 890 Mycobacterium, 547 Paragonimus, 2487 parasitology, 2294, 2299, 2305, 2329, 2331

n cxli

Pneumocystis, 2020 screening specimens, 284 specimen collection, transport, and handling, 278, 281, 298, 1415 expectorated, 278 induced, 278 viral infections, 1415 SRGA (Swedish Reference Group for Antibiotics), 1268–1269 St. Anthony’s fire, 2190 St. Louis encephalitis virus, 1645, 1648, 1650 St. Louis polyomavirus (STLPyV), 1803, 1805, 1810 ST EIA assay, 695 ST-246, 1831 Stability, definition, 132 Stable flies, 2513 Stachybotrys, 2190 Stachybotrys chartarum sick building syndrome, 2192 trichothecenes, 2190 Staffing models, for clinical microbiology laboratory, 45–46 Staggers, 2516 Stains, 321–323; see also specific stains blood films for parasites, 2334–2335 fungi, 1970 mycology, 1956–1959 parasitology, 2312–2314, 2316 stool samples for parasites, 2318–2319 virology, 1423–1425 Staminipila (kingdom), 1939 Standard fluid medium 10B (Shepard’s M10 medium), 344 Staphaurex Plus, 363 StaphPlex panel, 361, 364 Staphylinid beetle, 2521 Staphylococcaceae family, 354–372 antimicrobial susceptibilities, 368–371 clinical significance, 357, 360–361 collection, transport, and storage of specimens, 361 description of family, 354–356 differentiation of species, 355, 358–359 direct examination, 361–362 epidemiology and transmission, 356–357 evaluation, interpretation, and reporting of results, 371–372 identification, 362–366 isolation procedures, 362 serologic tests, 368 taxonomy, 354 typing systems, 367–368 Staphylococcal chromosomal cassette mec (SCCmec), 139, 1222, 1380 Staphylococcal enterotoxin B (SEB), 223 Staphylococcal scalded skin syndrome, 360, 365–366 Staphylococcal toxic shock syndrome, 360, 365 Staphylococci, see Staphylococcus Staphylococcus, 354–372 antibiotic resistance, 1214, 1216–1218, 1220–1223, 1229–1232, 1234–1235 automated detection, 1278–1279 inducible clindamycin resistance, 1290, 1295–1297 linezolid resistance, 1278 oxacillin resistance, 1278, 1289–1294 penicillin resistance, 1278, 1289 phenotypic methods for detecting, 1287, 1289–1297 vancomycin resistance, 1290, 1294– 1295

cxlii

n

SUBJECT INDEX

Staphylococcus (continued) antimicrobial susceptibilities, 368–371, 1174–1175, 1177–1182, 1184– 1191, 1193, 1195–1197 antimicrobial susceptibility testing, 1249– 1250, 1253–1256, 1259–1261, 1267, 1270 β-lactamase, 1302 β-lactamase tests, 1302–1303 clindamycin resistance detection, 1290, 1295–1297 D-zone test, 1267, 1296–1297 quality control and quality assessment, 1296 reporting of results, 1296 single-well broth dilution method, 1267, 1296 clinical significance, 357, 361 coagulase-negative (CoNS), 355–356 antibiotic resistance, 1230, 1249, 1278, 1287, 1289, 1292–1293, 1294 antimicrobial susceptibilities, 369–371, 1184, 1189–1190, 1197 antimicrobial susceptibility testing, 1254, 1259–1260, 1267, 1277 blood culture, 18 clinical significance, 360–361 epidemiology and transmission, 357 evaluation, interpretation, and reporting of results, 371–372 identification, 362–365 isolation procedures, 362 methicillin-resistant, 361, 1381 oxacillin resistance detection, 1278, 1292–1293 typing systems, 368 vancomycin-intermediate (VISS), 1294 coagulase-positive, 355, 360 epidemiology and transmission, 356–357 collection, transport, and storage of specimens, 361 commercial sources of chromogenic agar media for, 326 description, 354–356 differentiation of species, 355, 358–359 direct examination, 361–362 microscopy, 361 nucleic acid detection, 361–362 epidemiology and transmission, 356–357 evaluation, interpretation, and reporting of results, 371–372 on German cockroach body surface, 2513 identification, 362–366 agglutination assays, 363–364 by biochemical procedures, 364 by nucleic acid-based approaches, 365 by spectroscopic approaches, 365 by susceptibility tests, 364 coagulase production, 362–363 colony morphology, 362–363 of toxin-mediated syndromes, 365–366 in skin microbiome, 232 isolation procedures, 362 NaCl agar for, 341 oxacillin resistance detection, 1289–1294 by PCR, 1293–1294 chromogenic agars, 1293 in coagulase-negative staphylococci, 1292–1293 in S. aureus and S. lugdunensis, 1289– 1292 rapid tests, 1293 reporting results of tests, 1294

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 142

phenotypic methods for detecting antibacterial resistance, 1287, 1289–1297 serologic tests, 368 taxonomy, 354 typing systems, 367–368 vancomycin resistance detection, 1294– 1295 BHI-V6 screening agar test method, 1295 limitations of methods, 1295 quality control, 1295 reporting of results, 1295 Staphylococcus agar no. 110, 344 Staphylococcus agnetis, 357–358, 360 Staphylococcus arlettae, 355, 357–358 Staphylococcus aureus, 354–372 antibiotic resistance, 1214, 1216–1218, 1220–1223, 1229–1232, 1234–1235 automated detection, 1278–1279 clindamycin resistance, 1296 mupirocin resistance, 1290, 1297 oxacillin resistance detection, 1289– 1292 vancomycin resistance detection, 1290, 1294–1295 antimicrobial susceptibilities, 368–371, 1172, 1174–1175, 1181, 1184, 1186, 1188, 1195–1197 antimicrobial susceptibility testing, 1249– 1250, 1259–1261, 1267, 1277 as ESKAPE pathogen, 714 β-lactamase tests, 1302–1303 biothreat agent, 223 blood culture, 18, 24, 1293 BORSA (borderline oxacillin-resistant S. aureus), 1291–1292 CAMP test, 392, 481 characteristics of species, 355, 358 clinical significance, 357, 360 disinfectant resistance, 195 epidemiology and transmission, 356–357 epiglottitis, 301 evaluation, interpretation, and reporting of results, 371–372 GISA (glycopeptide-intermediate S. aureus), 369–370 identification, 362–366 in cystic fibrosis patients, 299 isolation procedures, 362 mecA gene during long-term preservation, 162 MRSA (methicillin-resistant S. aureus) antibiotic resistance, 1214, 1222, 1230– 1231, 1234–1235, 1289–1292, 1380–1382 antimicrobial susceptibilities, 368–371, 1173–1175, 1179, 1185, 1189– 1190, 1192, 1197 antimicrobial susceptibility testing, 1254 chromogenic agars for, 1293 clinical significance, 357 community-acquired (CA-MRSA), 1291 detection in blood, 24, 1293 epidemiology and transmission, 356–357 evaluation, interpretation, and reporting of results, 371–372, 1294 isolation procedures, 362 molecular detection, 1380–1382 nucleic acid detection, 361–362 oxacillin resistance detection, 1289– 1294 PCR detection, 1293–1294

rapid detection, 1249, 1293 specimen collection, transport, and handling, 302–303 transmission, 1214 treatment, 370–371 typing, 367–368 MSSA (methicillin-susceptible S. aureus) antimicrobial susceptibilities, 1174– 1175, 1185, 1189–1190, 1197 epidemiology and transmission, 356–357 molecular detection, 1380–1382 treatment, 370 nasal and nasopharynx carriage rate, 231 nucleic acid detection, 361–362 oxacillin resistance detection, 1289–1292 cefoxitin as surrogate for oxacillin, 1291 chromogenic agars, 1293 limitations of methods, 1292 oxacillin-salt agar screening test, 1291 PCR, 1293–1294 quality control, 1291 rapid tests, 1293 penicillin zone edge test, 1302 reference strains, 1264, 1267, 1291, 1296– 1297 serologic tests, 368 small-colony variants, 360, 370–371 specimen collection, transport, and handling, 300, 302–303, 361 staphylococcal enterotoxin B (SEB), 223 typing systems, 367–368 vancomycin resistance detection, 1294– 1295 BHI-V6 screening agar test method, 1295 limitations of methods, 1295 quality control, 1295 reporting of results, 1295 VISA (vancomycin-intermediate S. aureus), 369–370, 1230 antimicrobial susceptibilities, 1175, 1188–1189 automated system detection of, 1279 detection of, 1266, 1279, 1294–1295 disk diffusion test for, 1266 hVISA (heterogeneous VISA), 1230, 1295 reporting of, 1295 VRSA (vancomycin-resistant S. aureus) antimicrobial susceptibilities, 1175, 1189 automated system detection of, 1279 detection of, 1279, 1294–1295 reporting of, 1295 resistance mechanisms, 1230 Staphylococcus aureus ID, 343 Staphylococcus aureus subsp. anaerobius, 354– 355, 357–358, 360, 362, 909 Staphylococcus aureus subsp. aureus, 356, 363 Staphylococcus auricularis, 355–358, 362 Staphylococcus capitis, 355, 357, 360 Staphylococcus capitis subsp. capitis, 356, 358 Staphylococcus capitis subsp. urealyticus, 356, 358 Staphylococcus caprae, 3, 356 Staphylococcus carnosus, 354, 360 Staphylococcus carnosus subsp. carnosus, 357– 358 Staphylococcus carnosus subsp. utilis, 357–358 Staphylococcus caseolyticus, see Macrococcus caseolyticus Staphylococcus (Micrococcus) caseolyticus, see Macrococcus caseolyticus Staphylococcus chromogenes, 357, 359–360, 362, 364

SUBJECT INDEX Staphylococcus cohnii, 356, 360 Staphylococcus cohnii subsp. cohnii, 356, 358 Staphylococcus cohnii subsp. urealyticus, 356, 358 Staphylococcus condimenti, 357, 360 Staphylococcus delphini, 357, 359–360 Staphylococcus devriesei, 357–358, 362 Staphylococcus epidermidis anaerobic, 909 antimicrobial susceptibilities, 369, 1188, 1193, 1195, 1278, 1292–1293 blood culture, 24 clinical significance, 360 description, 354–355 epidemiology and transmission, 356–357 identification, 363–365 molecular detection of antibiotic resistance, 1381 Staphylococcus equorum, 355, 362 Staphylococcus equorum subsp. equorum, 357– 358 Staphylococcus equorum subsp. linens, 357– 358, 360 Staphylococcus felis, 357–358 Staphylococcus fleurettii, 354, 357–358 Staphylococcus gallinarum, 357–358 Staphylococcus haemolyticus, 354–358, 360, 362–363, 369, 1230, 1278, 1292 Staphylococcus hominis, 355, 357, 360, 363– 364, 1278, 1292 Staphylococcus hominis subsp. hominis, 356, 358 Staphylococcus hominis subsp. novobiosepticus, 356, 358 Staphylococcus hyicus, 357–358, 360, 363–364 Staphylococcus intermedius, 355, 357–358, 360, 363, 369 Staphylococcus kloosii, 356–358 Staphylococcus lentus, 354–355, 357–358, 362 Staphylococcus lugdunensis, 352, 354, 355, 356, 358, 360, 362–364, 369–371 blood culture, 18 molecular detection of antibiotic resistance, 1381 optochin-resistant strains, 352 oxacillin resistance, 1278, 1289, 1291 vancomycin resistance, 1294 Staphylococcus lutrae, 357–358 Staphylococcus massiliensis, 357–358 Staphylococcus microti, 357–358 Staphylococcus muscae, 357–358 Staphylococcus nepalensis, 357–358 Staphylococcus pasteuri, 356, 359–360 Staphylococcus petrasii, 357 Staphylococcus pettenkoferi, 356–357, 359–360 Staphylococcus piscifermentans, 357, 359–360 Staphylococcus pseudintermedius, 354, 357, 359–360, 363, 369, 371 Staphylococcus pulvereri, 357 Staphylococcus rostri, 357, 359 Staphylococcus saccharolyticus, 354–356, 359, 362, 909–910 Staphylococcus saprophyticus, 352, 355–356, 360, 363–364, 369, 371, 1195–1196, 1292 Staphylococcus saprophyticus subsp. bovis, 357, 359–360 Staphylococcus saprophyticus subsp. saprophyticus, 356–357, 359–360, 363 Staphylococcus schleiferi subsp. coagulans, 357, 359–360 Staphylococcus schleiferi subsp. rodentium, 359 Staphylococcus schleiferi subsp. schleiferi, 356, 359, 363

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 143

Staphylococcus sciuri, 362, 368, 1221, 1292 Staphylococcus sciuri subsp. carnaticus, 357, 359 Staphylococcus sciuri subsp. rodentium, 359 Staphylococcus sciuri subsp. sciuri, 359 Staphylococcus simiae, 356, 359 Staphylococcus simulans, 356, 359, 1292–1293 Staphylococcus stepanovicii, 3, 354, 357 Staphylococcus succinus subsp. casei, 359 Staphylococcus succinus subsp. succinus, 357, 359 Staphylococcus vitulinus, 354–355, 357, 359, 362, 368 Staphylococcus warneri, 355–356, 359–360, 362, 364 Staphylococcus xylosus, 356, 357, 359–360, 362, 368 Staphylococcus-Streptococcus selective medium, 344 Staphytect Plus, 363 Star of David appearance, 1619, 1623 Starch hydrolysis, 616 STAT! Campy assay, 1002 Stavudine adenoviruses, 1777 human immunodeficiency virus (HIV), 1870, 1873 Stavudine resistance, 1896–1898 Steam sterilization, 203–204 Steatorrhea, Cystoisospora belli and, 2428 STEC, see Escherichia coli, Shiga toxinproducing Stellantchasmus, 2482 Stenotrophomonas maltophilia, 615, 626–627 antibiotic resistance, 1226 antimicrobial susceptibilities, 804, 1176– 1178, 1180, 1186–1187, 1192– 1193 antimicrobial susceptibility testing, 1253, 1255–1256, 1270 carbapenemases, 1300 characteristics of, 802 clinical significance, 794–795 direct examination, 795 epidemiology and transmission, 793 identification, 801–802 isolation procedures, 797 taxonomy, 792 typing systems, 803 Sterilant, 190 Sterilization, 202–205 definition, 189, 202 ethylene oxide gas, 204 flash, 204 liquid, 205 monitoring, 203 ozone, 205 packaging, loading, and storage, 203 plasma, 204 principles, 202–203 steam, 203–204 Sterilox, 195 Steris system 1, 205 Steroids, Epstein-Barr virus and, 1739 Sterrad 100 sterilizer, 204 Sterrad NX system, 204 Stevens-Johnson syndrome, 1192, 1360, 1521 Stichodora, 2482 “Stick and ball” structures, 1806, 1808 Stick Crypto, 2441 Stick Crypto-Giardia, 2441 Sticktight flea, 2509 Stillbirth, syphilitic, 1061

n cxliii

Stinging and biting arthropods, 2518–2521 centipedes and millipedes, 2520 Hymenoptera, 2518 scorpions, 2519–2520 spiders, 2520–2521 urticating caterpillars, 2518–2519 STLVs (simian T-cell lymphotropic viruses), 1458–1459 Stokes method, 1269 Stomatitis Kingella, 655 measles, 1521 Treponema, 1061 Stomatococcus, 354, 1184 Stomatococcus mucilaginosus, 494; see also Rothia mucilaginosa Stomoxys calcitrans, 2519 Stool antigen test (SAT), for Helicobacter, 1018–1019, 1023 Stool specimen fungi, 1946, 1950–1951 Mycobacterium, 548 parasitology, 2293–2304, 2317–2324 commercial kits for immunodetection in samples, 2295 concentration wet mount, 2317–2318 culture of larval-stage nematodes, 2321– 2323 direct wet mount in saline, 2317 egg identification, 2320 hatching of schistosome eggs, 2323 helminth recovery and identification techniques, 2323 immunoassay methods, 2319–2320, 2322 key to identification of intestinal amebae, 2321 key to identification of intestinal flagellates, 2322 molecular methods, 2320–2321 permanent stained smears, 2318–2319 processing liquid stool, 2318 processing preserved stool, 2318–2319 specimen collection and processing options, 2301 stool collection, 2293–2294, 2296 stool preservation, 2300–2304 taenia solices, search for, 2324 test ordering, 2302 worm burden estimation, 2323 screening specimens, 284 viruses, 1415 Stool specimen preservation, 2300–2304 formalin, 2301–2303 modified polyvinyl alcohol, 2303–2304 polyvinyl alcohol (PVA), 2303–2304 Schaudinn’s fluid, 2303–2304 single-vial collection systems, 2303–2304 sodium acetate-acetric acid-formalin (SAF), 2302–2303 Storage of microorganisms, 113, 161–167 disaster preparedness, 167 freeze-drying (lyophilization), 164–165 cryoprotective agents, 165 methods, 165 preparation of microbes, 165 reconstitution, 165 storage, 165 storage vials, 165 long-term preservation methods, 162 freeze-drying (lyophilization), 164–165 newer technologies, 165 ultralow-temperature freezing, 162–164 procedures for specific organisms, 163, 166–167

cxliv

n

SUBJECT INDEX

Storage of microorganisms (continued) bacteria, 166 protozoa, 166 viruses, 167 yeasts and filamentous fungi, 166–167 short-term preservation methods, 161–162 direct transfer to subculture, 161 drying, 162 freezing at –20°C, 162 frequency of transfer, 161–162 immersion in oil, 162 maintenance medium, 161 quality control procedures, 162 storage conditions, 161 storage in distilled water, 162 ultralow-temperature freezing, 162–164 cryoprotective agents, 163–164 freezing method, 164 preparation of microbes for freezing, 164 specialized storage systems, 164 storage vials, 163 thawing, 164 virus samples, 1410–1412 Storage vials for freeze-drying, 165 for ultralow-temperature freezing, 163 Strain, definition, 132 Strain catalogues, 150–151 Stramenopila (kingdom), 1936, 2200, 2203 Strand displacement amplification, 64–66, 1110–1112 Stratify JCV test, 1810, 1812 Strep B carrot broth, 344 Streptobacillus antimicrobial susceptibilities, 662 clinical significance, 655 direct examination, 656 epidemiology and transmission, 654 identification, 660–661 isolation procedures, 658 specimen management, 283 taxonomy and description of, 653 Streptobacillus moniliformis, 653, 655–662 Streptococcal toxic shock syndrome (STSS), 385–386 Streptococci, see Streptococcus Streptococcus, 355, 383–397 alpha-hemolytic streptococci, colony morphology of, 390–391 antibiotic resistance, 1220, 1279, 1319– 1320 automated detection, 1279 clindamycin resistance, 1297–1298 molecular detection, 1383 phenotypic methods for detecting, 1297–1298 antigen detection in CSF, 388 S. agalactiae in urogenital tract samples, 388 S. pneumoniae in urine samples, 388 S. pyogenes from throat specimen, 388 antimicrobial susceptibilities, 396–397, 1173–1175, 1177, 1179, 1181– 1182, 1184–1191, 1195–1197 beta-hemolytic streptococci, 396–397, 1279 S. pneumoniae and S. viridans group streptococci, 397 antimicrobial susceptibility testing, 1250, 1265, 1267, 1270, 1319–1320 commercial test methods, 1320 incidence of resistance, 1319–1320 reference test methods, 1320

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 144

strategies for testing and reporting of results, 1320 beta-hemolytic antimicrobial susceptibilities, 396–397 antimicrobial susceptibility testing, 1319–1320 colony morphology, 390 identification by Lancefield antigen immunoassays, 391 identification with phenotypic tests and MALDI-TOF MS, 391–392 phenotypic characteristics of, 384 rectal swab screening for, 303 Streptococcus pyogenes, 389 clindamycin resistance detection D-zone test, 1267, 1297–1298 quality control and quality assessment, 1298 reporting of results, 1298 single-well broth dilution method, 1267, 1298 clinical significance, 385–387 collection, transport, and storage of specimens, 387 description of genus, 384–385 direct examination, 387–389 antigen detection, 388 microscopy, 387–388 epidemiology and transmission, 385 evaluation, interpretation, and reporting of results, 397 identification, 390–395, 426 colony description, 390–391 of beta-hemolytic streptococci, 391–392 of S. pneumoniae and S. viridans group streptococci, 392–395 in esophageal microbiome, 229 in gastric microbiome, 230 in small intestinal microbiome, 230 isolation procedures, 389–390 phenotypic methods for detecting antibacterial resistance, 1297–1298 serologic tests, 396 taxonomy, 383–384 typing systems, 395–396 viridans, 1177, 1181, 1184–1185, 1187, 1197 antibiotic resistance, 1222 antimicrobial susceptibilities, 397, 1172, 1177, 1187 antimicrobial susceptibility testing, 1319–1320 identification, 392–395 taxonomy, 383–384 Streptococcus adjacens, 422 Streptococcus agalactiae, 383 antibiotic resistance, 1230, 1279, 1319– 1320 antigen detection of S. agalactiae in urogenital tract samples, 388 antimicrobial susceptibilities, 396–397, 1197 antimicrobial susceptibility testing, 1277, 1319–1320 CAMP test, 392 clinical significance, 386 collection, transport, and storage of specimens, 387 colony morphology, 390 detection in CSF, 388 epidemiology and transmission, 385 identification, 391–392 by Lancefield antigen immunoassays, 391

isolation procedures, 389–390 nucleic acid detection techniques, 388– 389 phenotypic characteristics, 384 taxonomy, 383 typing, 396 Streptococcus alactolyticus, 384, 387, 393–394 Streptococcus anginosus group, 352 antibiotic resistance, 1230, 1319–1320, 1352 clinical significance, 386–387 colony morphology, 390 evaluation, interpretation, and reporting of results, 397 identification, 391, 393 by Lancefield antigen immunoassays, 391 microscopy, 387 phenotypic characteristics, 384, 393, 394 taxonomy, 383–384 Streptococcus anginosus subsp. whileyi, 384 Streptococcus australis, 383, 386 Streptococcus bovis, 1172 Streptococcus bovis group, 352, 387 antibiotic resistance, 1319 colony morphology, 391 evaluation, interpretation, and reporting of results, 397 identification, 393–394 phenotypic characteristics, 393 taxonomy, 383–384 Streptococcus canis, 384, 385 Streptococcus constellatus, 384, 386–387, 393 Streptococcus constellatus subsp. constellatus, 393 Streptococcus constellatus subsp. pharyngis, 393 Streptococcus constellatus subsp. viborgensis, 384 Streptococcus criceti, 384, 387, 393 Streptococcus cristatus, 383, 386 Streptococcus defectivus, 422 Streptococcus devriesei, 384, 387, 393 Streptococcus didelphis, 385 Streptococcus downei, 384, 387, 393 Streptococcus dysgalactiae subsp. equisimilis, 396, 397 Streptococcus dysgalactiae subsp. dysgalactiae, 383, 384, 385 Streptococcus dysgalactiae subsp. equisimilis, 383, 385 clinical significance, 386 colony morphology, 390 phenotypic characteristics, 384 taxonomy, 383 Streptococcus equi subsp. equi, 384, 385, 386 Streptococcus equi subsp. zooepidemicus, 384, 385 Streptococcus equinus, 384, 387, 393–394 Streptococcus ferus, 384, 387, 393 Streptococcus gallolyticus, 384, 387, 393–394, 1320 Streptococcus gallolyticus subsp. gallolyticus, 394, 397 Streptococcus gallolyticus subsp. pasteurianus, 394 Streptococcus gordonii, 383, 384, 386 Streptococcus group B antibiotic resistance, 1222–1223 commercial sources of chromogenic agar media for, 326 Streptococcus hongkongensis, 387 Streptococcus hyovaginalis, 384, 387, 393 Streptococcus infantarius, 384, 387, 393–394 Streptococcus infantarius subsp. coli, 394

SUBJECT INDEX Streptococcus infantis, 383, 386 Streptococcus iniae, 387, 391 Streptococcus intermedius, 384, 386–387, 393 Streptococcus lactarius, 383 Streptococcus macacae, 384, 387, 393 Streptococcus massiliensis, 383, 386 Streptococcus milleri, 384, 1319 Streptococcus milleri group, 390 Streptococcus mitis, 386, 1381 antibiotic resistance, 1222, 1320 endocarditis, 229 evaluation, interpretation, and reporting of results, 397 identification, 394 Streptococcus mitis group antibiotic resistance, 1319 clinical significance, 386 identification, 393 phenotypic characteristics, 393 taxonomy, 383–384 Streptococcus morbillorum, 422 Streptococcus mutans, 384 Streptococcus mutans group antibiotic resistance, 1319 clinical significance, 387 identification, 393 phenotypic characteristics, 393 taxonomy, 383–384 Streptococcus oligofermentans, 383, 386 Streptococcus oralis, 229, 383, 386, 393, 1320 Streptococcus orisratti, 383, 386 Streptococcus parasanguinis, 383, 386 Streptococcus parvulus, see Atopobium parvulum Streptococcus peroris, 383, 386 Streptococcus pneumoniae antibiotic resistance, 1212, 1216–1218, 1222, 1228, 1230–1231, 1234– 1235, 1279, 1383 β-lactam-resistant, 1383 cephalosporin resistance, 1316 clindamycin resistance, 1297–1298, 1316 erythromycin resistance, 1316 fluoroquinolone resistance, 1319 incidence, 1315–1316, 1319 macrolide resistance, 1316 MLS-type resistance, 1316 penicillin resistance, 1315–1316 trimethoprim-sulfamethoxazole resistance, 1319 antigen detection of S. pneumoniae in urine samples, 388 antimicrobial susceptibilities, 397, 1172, 1174, 1179, 1182, 1184, 1186, 1192–1193, 1195, 1197 antimicrobial susceptibility testing, 1250, 1253, 1267, 1277, 1315–1316, 1319 commercial methods of testing, 1319 incidence of resistance, 1315–1316, 1319 reference test methods, 1319 strategies for testing and reporting of results, 1319 blood culture, 18 clinical significance, 386 colony morphology, 390–391 detection in blood, 20 detection in CSF, 388 epidemiology and transmission, 385 epiglottitis, 301 evaluation, interpretation, and reporting of results, 397

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 145

identification, 391, 392–395 microscopy, 387 nucleic acid detection techniques, 389 optochin susceptibility, 394 optochin-resistant strains, 352 reference strains, 1298, 1315 Trypticase soy agar, with sheep blood and gentamicin for, 346 Trypticase soy agar, with sheep blood, sucrose, and tetracycline for, 346 typing, 395–396 vaccine, 386 Streptococcus porcinus, 384, 387, 391 Streptococcus pseudopneumoniae, 383, 386 Streptococcus pseudoporcinus, 391 Streptococcus pyogenes antibiotic resistance, 1231, 1319–1320 antimicrobial susceptibilities, 396, 1172, 1174, 1186, 1195, 1197 antimicrobial susceptibility testing, 1319– 1320 bacitracin susceptibility, 391 clinical significance, 385–386 colony morphology, 390–391 direct antigen detection of S. pyogenes from throat specimens, 388 epidemiology and transmission, 385 epiglottitis, 301 evaluation, interpretation, and reporting of results, 397 nucleic acid detection techniques, 388 phenotypic characteristics, 384 specimen collection, transport, and handling, 299–300, 387 throat cultures, 389 typing, 396 Streptococcus ratti, 384, 387, 393 Streptococcus salivarius, 393 Streptococcus salivarius group antibiotic resistance, 1319 clinical significance, 387 colony morphology, 391 identification, 393 phenotypic characteristics, 393 taxonomy, 383–384 Streptococcus sanguinis, 383, 386, 1320 Streptococcus selective medium, 344 Streptococcus sinensis, 383, 386 Streptococcus sobrinus, 384, 393 Streptococcus suis, 385, 387 Streptococcus thermophilus, 384, 393 Streptococcus tigurinus, 383 Streptococcus uberis, 384 Streptococcus vestibularis, 384, 393 Streptodornase, 231 Streptogramin(s), 1189–1190 adverse effects, 1190 mechanism of action, 1189 pharmacokinetics, 1190 spectrum of activity, 1190 Streptogramin resistance, 1189, 1215, 1231 Streptomyces aminoglycoside production, 1181 aminoglycoside-modifying enzymes, 1220 clinical significance, 519 description of genus, 511 fosfomycin production, 1196 identification, 438, 522, 524 isolation procedures, 521 microscopy, 520–521 morphologic characteristics, 508 streptogramin production, 1189 taxonomy, 505 Streptomyces albus, 514, 519

n cxlv

Streptomyces avermitilis, 2533 Streptomyces bikiniensis, 514, 519 Streptomyces cinereoruber, 514 Streptomyces erythreus, 1182, 1231 Streptomyces griseus, 510, 514 Streptomyces lincolnensis, 1185 Streptomyces mediterranei, 1194 Streptomyces mutabilis subsp. capreolus, 1360 Streptomyces nodosus, 2228 Streptomyces orientalis, 1187 Streptomyces roseosporus, 1187 Streptomyces somaliensis, 513, 519 Streptomyces sudanensis, 514 Streptomyces thermovulgaris, 514, 519 Streptomyces venezuelae, 1193 Streptomycin, 1180–1182, 1199 antimicrobial susceptibility testing, 1255, 1260, 1365, 1367 for Mycobacterium infection, 1358–1360 high-level streptomycin resistance, 1278 Streptomycin resistance Mycobacterium tuberculosis complex, 1360 Yersinia pestis, 1324 Streptosel agar, 345 Stress adaptation, azole resistance and, 2243 Strobila, 2471, 2476 Stroma, 1942 Strongylida (order), 2289, 2454 Strongyloidea (superfamily), 2289 Strongyloides, 2322, 2326, 2448, 2459 Strongyloides fuelleborni, 2458 Strongyloides IgG ELISA, 2458 Strongyloides stercoralis, 2456–2458 agar plate culture for, 2321, 2457 antiparasitic agent resistance, 2555 clinical significance, 2457 commercial kits for immunodetection of serum antibodies, 2296 description, transmission, and life cycle, 2454, 2456–2457 eggs, 2456 larvae, 2452, 2456 worms, 2456–2457 detection, 2321–2323, 2325, 2329, 2331, 2457, 2459 diagnosis, 2457–2458 epidemiology and prevention, 2457 sputum specimen, 2305 taxonomy, 2456 treatment, 2455, 2458, 2531, 2534 Strongyloidiasis, 2296, 2448, 2457 Strongyloididae (family), 2289 Stuart transport medium, modified, 345 Subacute sclerosing panencephalitis (SSPE), 1520 Subculture, 161, 166 Subcutaneous infection/lesions Basidiobolus ranarum, 2099 chromoblastomycosis, 2161 Coccidioides, 2114 dirofilariasis, 2499 hyaline fungi, 2075–2076 Lagenidium, 2198, 2204 Onchocerca volvulus, 2466 phaeohyphomycoses, 2161–2163 Pythium insidiosum, 2201 sporotrichosis, 2164 Substage condenser, 8 Subtype, definition, 132 Subtyping methods, see also specific organisms amplified fragment length polymorphism, 137–138 binary typing, 143 characteristics, 131–135

cxlvi

n

SUBJECT INDEX

Subtyping methods (continued) convenience parameters, 134–135 discriminatory power, 134 reproducibility and stability, 132, 134 typeability, 131 data interpretation, 149–150 DNA microanalysis, 144–145 forensic microbiology, 148 k-mer analysis, 143–144 mass spectrometry, 145 MLST, 143 molecular surveillance, 145–148 cluster detection/outbreak investigations, 145–146 molecular serotyping, 145 source attribution, 146–147 non-target-specific methods, 135–137 IS6110 fingerprinting, 136–137 PCR ribotyping, 137 PCR-based, 137 plasmid profiling, 135–136 pulsed-field gel electrophoresis, 136 random amplification of polymorphic DNA/arbitrarily primed PCR, 137 repetitive element PCR, 137 restriction fragment length polymorphisms, 136 whole-genome mapping, 137 selection of method, 148–149 single nucleotide polymorphism (SNP)base methods, 141–143 target-specific method, 138–141 clustered regularly interspaced short palindromic repeats (CRISPs) analysis, 140–141 gene sequencing, 139–140 PCR-RFLP, 138–139 variable-number tandem-repeat analysis, 140 whole-genome sequencing, 141 validation of method, 149 Succinatimonas, 969 Succinatimonas hippei, 969 Succinivibrionaceae (family), 969 Sucrose, as cryoprotectant in transport medium, 1409 Sucrose-phosphate-glutamate containing 1% bovine albumin (SPGA), 167 Sucrose-teepol-tellurite agar, 345 Sudan ebolavirus, 1670 Sudan virus, 1670, 1672–1673, 1677–1678 Sulbactam, 1177–1178 with ampicillin, 1177–1178, 1198 Sulfa drugs, for Acanthamoeba, 2395 Sulfadiazine, 1191, 1199 Acanthamoeba, 2394 Balamuthia mandrillaris, 2395 Toxoplasma gondii, 2381–2382 Sulfadoxine, 1199 Sulfadoxine-pyrimethamine, 2349, 2540– 2541, 2564 Sulfadoxine-pyrimethamine resistance, 2550– 2553 Sulfamethizole, 1199 Sulfamethoxazole, 1199 Sulfanilamide, 1191 Sulfanilic acid, 319 Sulfide-indole-motility medium, 345 Sulfite reductase activity, 615 Sulfonamide(s), 1191–1192 adverse effects, 1192 antimicrobial susceptibility testing, 1256, 1261

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 146

concentration in serum, 1199 mechanism of action, 1191 pharmacology, 1191 spectrum of activity, 1191–1192 Toxoplasma gondii, 2381–2382 with trimethoprim, 1191–1192, 1199 Sulfonamide resistance, 1234–1235 acquired resistance to sulfonamides, 1234– 1235 common associations of resistance mechanisms, 1215 intrinsic resistance, 1234 Neisseria meningitidis, 1324 Sulfur granules, 925 Sulfuric acid method, of digestion and decontamination, 560 Sulfurospirillum, 998 Sulfurospirillum deleyianum, 998 Super E-Mix, 1426, 1543 coxsackie B virus detection in, 1427 virus susceptibility profiles, 1429 Superficial mycoses, 2146–2149 black piedra, 2148 phaeohyphomycoses, 2161–2162 tinea nigra, 2147–2148 tinea versicolor, 2146–2147 white piedra, 2148–2149 Superficial wound screening specimens, 284 specimen collection, 271 Superoxidized water, 195 Suramin, 2544, 2551, 2555 adverse effects, 2544 African trypanosomiasis, 2564 mechanism of action, 2544 pharmacokinetics, 2544 spectrum of activity, 2544 Trypanosoma brucei, 2367 Sure-Vue RSV, 1504 Surface-enhanced laser desorption ionization-TOF MS, 263 Surfaces, cleaning and disinfecting, 196–197 Surfacine, 195 Surgical hand disinfection (rub-in), 187 Surgical hand washing (scrub), 187 Surgical wound infections, Mycobacterium, 600 Surveillance cultures from patients, hospital personnel, and the environment, 113 disease, 120–121 Susceptible dose-dependent (SDD), 2257 Sutterella characteristics of genus, 970–971 clinical significance, 974 identification, 977, 981 taxonomy, 969 Sutterella parvirubra, 969, 981 Sutterella wadsworthensis, 969, 974, 977, 981, 994, 997, 1348 Sutterellaceae (family), 969 Suttonella antimicrobial susceptibilities, 662 clinical significance, 655 direct examination, 656 identification, 661 isolation procedures, 658 taxonomy and description of, 653 Suttonella indologenes, 653, 655–656, 658, 661–662 Swabs for specimen collection, 270–271 initial sample handling, 285 Swedish Reference Group for Antibiotics, 1268–1269

Swimmer’s itch, 2480, 2486 SWIN (Sensititre Windows software) data management system, 34, 1275, 1277 Swine, influenza viruses, 1471 Swine erysipelas, 468 Swollen-baby syndrome, 1673 Sydowia polyspora, 2155 Symbiogenesis, 2286 Sympodial, 1940, 1942 Syngamidae (family), 2289 Synanamorph, 1942, 2058, 2154–2155 Syncephalastraceae (family), 2088, 2096 Syncephalastrum, 2088, 2096 Syncephalastrum racemosum, 2088, 2096, 2101 Synergies plus panel, 364, 1276 Synergistetes (phylum), 967–968, 974, 980, 983 Synnema, 1942, 2058 Synovial fluid specimen collection, transport, and storage guidelines, 276 fungi, 1946–1947, 1950 Synovitis Dolosigranulum pigrum, 424 Methylobacterium, 830 Mycobacterium tuberculosis, 538 Syphicheck-WB, 1069 Syphilis, 1055–1075 antimicrobial susceptibilities, 1072, 1172 clinical significance, 1059–1061 collection, transport, and storage of specimens, 1061–1063 congenital clinical significance, 1059 criteria for diagnosis, 1061 tests for, 1062, 1075 description of agents, 1055–1057 direct examination, 1063 endemic treponematoses, 1055–1056, 1058, 1061 epidemiology and transmission, 1057–1058 evaluation, interpretation, and reporting of results, 1072–1075 direct detection of Treponema pallidum, 1072–1073 serologic tests, 1073–1074 syphilis tests in HIV infection, 1075 tests for congenital syphilis, 1075 tests for neurosyphilis, 1074 HIV and, 1058–1059, 1073, 1075 isolation procedures for Treponema pallidum, 1064 natural course of untreated, 1059 neurosyphilis clinical significance, 1059 criteria for diagnosis, 1060–1061 tests for, 1062, 1074 rabbit infectivity testing (RIT), 1055 stages/manifestations, 1056, 1060–1061 treponemal tests for syphilis, 1066–1072 chemiluminescence immunoassays (CLIAs), 1071–1072 combined treponemal IgM/IgG EIAs, 1068 conventional, 1066, 1070, 1074 EIAs, 1067–1068, 1070–1071, 1074 FTA-ABS test, 1066, 1070, 1074 immunoblot assays, 1070, 1072 MHA-TP test, 1066, 1070, 1074 multiplex flow immunoassays, 1071– 1072 POC (point-of-contact), 1066, 1071– 1072, 1074

SUBJECT INDEX rapid combination nontreponemal and treponemal tests, 1070 rapid treponemal tests, 1069, 1071– 1072, 1074 TPHA, 1066, 1070 TP-PA test, 1066, 1070, 1074 treponemal IgM or IgG EIAs, 1067 typing systems for Treponema pallidum, 1065 venereal clinical significance, 1059–1061 epidemiology and transmission, 1057– 1058 Syphilis EIA II, 1068, 1071 Syphilis Fast, 1069 Syphilis Health Check, 1069 Syphilis Total Antibody, 1068 Syrphid, 2517 Systemic mycoses, dimorphic fungi causing, 2109–2123 T. pallidum hemagglutination assay (TPHA), 1066, 1070 T. pallidum particle agglutination assay (TPPA), 1066, 1070, 1074 T. phagedenis Reiter (Sorbent), 1070 T-2 toxin, 2189–2190, 2192 T2Candida test, 1979 T5000 biosensor technology, 263 T5000 Universal Biosensor/PLEX-ID, 72 Tacaribe virus, 1669, 1671 Tachycardia, scorpion venom and, 2520 Tachyzoites, Toxoplasma gondii, 2373–2374 Taenia, 2502 detection, 2320 eggs, 2449 taenia solices, search for, 2324 treatment, 2533 Taenia (genus), 2473–2474 Taenia asiatica, 2473 Taenia crassiceps, 2471, 2477 Taenia multiceps, 2471, 2477, 2502 Taenia saginata, 2471–2474 clinical significance, 2473 collection, transport, and storage of specimens, 2473 description, 2473 detection, 2320 direct examination, 2474 epidemiology, transmission, and prevention, 2473 evaluation, interpretation, and reporting of results, 2474 microscopy, 2474 nucleic acid detection, 2474 serologic tests, 2474 taxonomy, 2473 treatment, 2474, 2531 Taenia saginata asiatica, 2473 Taenia serialis, 2477 Taenia solium, 2471–2472, 2474–2475, 2487 clinical significance, 2474 collection, transport, and storage of specimens, 2474 commercial kits for immunodetection of serum antibodies, 2296 description, 2474 detection, 2320, 2328–2330 direct examination, 2475 epidemiology, transmission, and prevention, 2474 evaluation, interpretation, and reporting of results, 2475 microscopy, 2475

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 147

neurocysticercosis, 2474, 2476, 2477 nucleic acid detection, 2474 serologic tests, 2475 taxonomy, 2474 treatment, 2475, 2531–2532 Taenia taeniaeformis, 2502 Taeniasis, 2474 Taeniidae (family), 2291, 2473–2474 Tafenoquine, 2539 Tahyna virus, 1645 Taï Forest virus, 1670, 1672 Taiwan taenia, 2473 Talaromyces, 2045–2048 Talaromyces marneffei, 2045–2048, 2115 antifungal susceptibility testing, 2271 antimicrobial susceptibilities, 2048 clinical significance, 2046 colony morphology, 2047 culture, 1951 cycloheximide inhibition, 1951, 1955 direct examination, 2046 epidemiology and transmission, 2048 evaluation, interpretation, and reporting of results, 2049 identification, 2047 isolation, 2046–2047 microscopy, 1966, 1976, 2046 nucleic acid detection, 2046 serologic tests, 2047–2048 specimen collection, transport, and processing, 1947, 1949 taxonomy, 2045–2046, 2048 typing systems, 2047 Talaromyces piceus, 2048 Talaromyces purpurogenus, 2048 Talaromyces rugulosus, 2048 Talaromyces verruculosus, 2048 Tamiami virus, 1669, 1671 TAMRA, 1690 Tanapox virus, 1828–1829, 1831, 1837 Tannerella characteristics of genus, 970–971 clinical significance, 971–972 taxonomy, 967, 969 Tannerella forsythia, 229, 967, 971–972, 978 Tapeworms, see Cestodes Taphrina, 2015 Tarantulas, 2521 Target amplification techniques, 57–67; see also specific techniques digital PCR, 63, 64 helicase-dependent amplification (HDA), 66–67, 69 loop-mediated amplification (LAMP), 66, 68 multiplex PCR, 58–60 nested PCR, 58 PCR, 57, 59 real-time PCR, 60–63 reverse transcriptase PCR (RT-PCR), 57– 58 strand displacement amplification (SDA), 64–66 transcription-based amplification methods, 63–64, 65 Tataguine virus, 1645 Tatlockia, 887 Tattoo ink infection, Mycobacterium chelonae and, 596 Tatumella, 718, 721, 726 collection, transport, and storage of specimens, 722 description of genus, 715 Tau protein, 1864

n cxlvii

Taxonomy, 255–265; see also specific organisms classification of bacteria, 255–265 criteria for species delineation, 256 described, 255 genomic threshold for species definition, 257–258 identification and classification methods, 259–263 major groups of bacteria, 258–259 molecular, 131 multilocus sequence analysis, 257 nomenclature, 263–264 polyphasic species concept, 256–257 uncultured bacteria, 259 viruses, 1393–1402 Tazobactam, 1173, 1178 with piperacillin, 1178, 1199 TB database, 151 TCBS agar, 345 TechLab E. histolytica II kit, 2404 TechLab Giardia II kit, 2411 Tedizolid, 1190 Teicoplanin, 1187–1188, 1199, 1260 Teicoplanin resistance, 1229–1230 Telaprevir, for hepatitis C virus, 1601–1602, 1879, 1901 Telaprevir resistance, 1901–1902, 1917 Telavancin, 1187–1189, 1199 Telbivudine resistance, 1851, 1900, 1917, 1921 Telenti fragment, 602 Teleomorph, 1936–1938, 1942, 2153–2155 Telithromycin, 1184–1185 adverse effects, 1185 antimicrobial susceptibility testing, 1256 concentration in serum, 1199 mechanism of action, 1184 pharmacology, 1184 spectrum of activity, 1184 Telithromycin resistance, 1231 Tellurite reductase activity, in Corynebacterium, 481 TEM β-lactamases, 1223–1226, 1299, 1320, 1322, 1331 Temafloxacin, 1179 Temocillin, 1172 Tenofovir for hepatitis B virus, 1881–1882, 1900 for HIV, 1870, 1873 Tenofovir resistance, 1917 hepatitis B virus, 1851, 1900 HIV, 1896–1898 Tenosynovitis Mycobacterium kansasii, 542 Mycobacterium szulgai, 543 Terbinafine, 2223–2224 antifungal susceptibility testing, 2255– 2273 chromoblastomycosis, 2167 Conidiobolus lamprauges, 2102–2103 dermatophytes, 2145 eumycotic mycetoma fungi, 2181–2182 Fusarium, 2069 melanized fungi, 2167 Pythium insidiosum, 2203 scedosporiosis, 2167 spectrum of activity, 2224 Talaromyces marneffei, 2048 yeast species, MICs for, 2005 Terbinafine resistance, 2223, 2239 Terminal restriction fragment length polymorphism (TRFLP), 226, 229 Termites, 2522

cxlviii n

SUBJECT INDEX

Tetanospasmin, 947 Tetanus, 947–948 Tetany, scorpion venom and, 2520 Tetracycline(s), 1185–1187 adverse effects, 1187 antimicrobial susceptibility testing, 1256, 1260 concentration in serum, 1199 Dientamoeba fragilis, 2413 malaria, 2564 mechanism of action, 1186 pharmacology, 1186 spectrum of activity, 1186–1187 Tetracycline resistance, 1186, 1216, 1233– 1234 Arcanobacterium haemolyticum, 1328 Neisseria gonorrhoeae, 1322–1323 Tetragenococcus, 422 Tetragenococcus solitarius, 422 Tetraparvovirus (genus), 1818, 1824 Tetrathiobacter, 838 Tetrathiobacter kashmirensis, 632–633 Tetrathionate broth, Hajna, 345 Tetrazolium tolerance agar, 345 Thallic conidiogenesis, 1939, 1942 Thallus, 1935, 1937, 1942 Thamnidiaceae (family), 2087 Thawing frozen samples, 164 Thayer-Martin medium, 345 Thayer-Martin medium, modified, 345 Thecamoebidae (family), 2387 Theileriasis, 2330 Thelazia, 2328 Thelaziidae (family), 2289 Thelazioidea (superfamily), 2289 Thermo Remel Candida albicans test kit, 2002 Thermoactinomyces chemotaxonomic and lysosome growth characteristics, 509 description of genus, 511 identification, 438 morphologic characteristics, 508 Thermoactinomycetaceae, 441 Thermoascaceae (family), 2073 Thermoascus, 2062, 2069 Thermoascus crustaceus, 2062 Thermoascus taitungiacus, 2062 Thiabendazole Capillaria philippinensis, 2497 Dracunculus medinensis, 2497 Strongyloides stercoralis, 2458 Trichinella, 2495 Thick blood films parasites, 2306, 2333–2335 Plasmodium, 2341–2342, 2345 preparation, 2333 proper examination, 2335 staining, 2334–2335 Thielavia, 2071 Thin blood films parasites, 2306, 2333–2335 Plasmodium, 2341–2342, 2344–2346 preparation, 2333 proper examination, 2335 staining, 2334–2335 ThinPrep Pap test vials, 1414 Thioglycolate bile broth, 345 Thioglycolate medium, enriched, 345 Thogoto virus, 1645 Thorny-headed worms, see Acanthocephalans Thottapalayam virus, 1664 3M Rapid Detection RSV test, 1504

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 148

Throat infection, see also Pharyngitis; Sore throat anaerobic Gram-negative rods, 972 Throat specimen collection, transport, and handling, 279, 299–300 Gram stain and plating medium recommendations, 286 Thrombocytopenia arenaviruses, 1674 carbapenems, 1177 cephalosporins, 1175 chloramphenicol, 1193 Epstein-Barr virus, 1739 linezolid, 1191 measles, 1521 penicillins, 1173 rifampin, 1195 spider envenomation, 2520 sulfonamides, 1192 Trypanosoma lewisi, 2368 varicella-zoster virus, 1705 Thrombophlebitis Fusarium, 2058 macrolides, 1183 Thrush, Saccharomyces cerevisiae and, 1994 Thymidine kinase, 1712 herpes simplex virus (HSV), 1894–1895 varicella-zoster virus, 1895 Thyroiditis, human T-cell lymphotropic viruses (HTLVs) and, 1461 Thysanura (order), 2522 Ticarcillin, 1173 antimicrobial susceptibility testing, 1255, 1259 concentration in serum, 1199 with clavulanic acid, 1177, 1199 Ticarcillin-clavulanic acid antimicrobial susceptibility testing, 1255, 1259 Bacteroides fragilis group susceptibility, 1346 Tick paralysis, 2516 Tick-borne encephalitis, 125, 2523 Tick-borne encephalitis virus (TBEV), 125, 1644–1645, 1647, 1650 Tick-borne lymphadenopathy, 1125 Tick-borne relapsing fever, 1037–1041; see also Borrelia Ticks as vectors, 2507, 2511–2513, 2515 degree of engorgement, estimating, 2523 hypostome, 2512 keys to, 2514, 2522 reactivation of pathogens, 2521, 2523 Tigecycline adverse effects, 1187 antimicrobial susceptibility testing, 1261 concentration in serum, 1199 mechanism of action, 1186 pharmacology, 1186 spectrum of activity, 1186–1187 Tigecycline resistance, 1234, 1347 TIGRIS system, 74 Tilletiaceae (family), 2075 Time to positivity, 18 Tindazole, 2564, 2566 Tinea, 2135–2136 Tinea barbae, 2136 Tinea capitis, 2136 Tinea corporis, 2136 Tinea cruris, 2136 Tinea manuum, 2136–2137 Tinea nigra, 2147–2148, 2162

Tinea pedis, 2058, 2136–2137 Tinea unguium, 2136 Tinea versicolor, 2146–2147 Tinidazole, 1194, 1199 Entamoeba histolytica, 2405 Giardia duodenalis, 2412 Trichomonas vaginalis, 2415 Tinidazole resistance, 2551, 2553 Tinnitus, arenaviruses and, 1673 Tinsdale agar, 345, 481 Tipranavir, for human immunodeficiency virus (HIV), 1871, 1876–1877 Tipranavir resistance, 1897–1898 Tissue infection Arcanobacterium haemolyticum, 479 Mycobacterium genavense, 542 Mycobacterium kansasii, 542 Mycobacterium xenopi, 543 Photobacterium damselae, 765 Trueperella pyogenes, 479 Vibrio fluvialis, 765 Tissue specimen, see also specific tissues collection, transport, and handling, 271, 279, 281, 289–292 bone marrow, 291 cellulitis, 292 lymph nodes, 291–292 necrotizing fasciitis, 292 placenta, 292 quantitative culture, 292 uncultivable bacteria, 292 fungi, 1946–1948, 1951 Gram stain and plating medium recommendations, 286 initial sample handling, 285 Mycobacterium, 548 parasitology, 2306 viruses, 1415–1416 Tityus, 2520 tlyA gene, 1360 TMPD/DMPD, 319 T/NK lymphoma, Epstein-Barr virus and, 1739–1741 TNP-470, 2216 Tobramycin, 1181–1182, 1199 antimicrobial susceptibility testing, 1255, 1260 for Mycobacterium chelonae, 1359 Todd-Hewitt broth, 345 Togaviridae (family), 1399–1401, 1525, 1644 Tolerance, 1212–1213 Toluidine blue O, 1958 Toluidine blue stain, for fungi, 1970, 1975 Toluidine red unheated serum test (TRUST) assays, 1062, 1066 Tolumonas, 752 Tongue worms, 2516 Tonsillitis adenoviruses, 1771 Corynebacterium diphtheriae, 1327 Epstein-Barr virus, 1739 Fusobacterium, 973 Neisseria gonorrhoeae, 636 Prevotella, 973 Tonto Creek virus, 1669, 1672 Tooth disorders, tetracyclines and, 1187 Topoisomerase IV, 1179 Topoisomerase IV, mutations in, 1218, 1232 Topotecan polyomavirus, 1811 progressive multifocal leukoencephalopathy (PML), 1811 TORCH (toxoplasmosis, other, rubella, cytomegalovirus, and herpes simplex virus) panels, 1530

SUBJECT INDEX Torovirinae (subfamily), 1398, 1565 Torovirus (genus), 1398 Toscana virus, 1645, 1651 Total laboratory automation, 49–51 BD-Kiestra TLA concept, 49–50 bioMérieux Concept FMLA, 50 digital imaging, 51 WASP Lab, 50–51 TOTAL-FIX, 2311 Toxascaris, 2448 Toxic shock syndrome Finegoldia magna, 911 staphylococcal, 360, 365 Toxigenic culture, Clostridium difficile, 951 Toxin B gene, Clostridium difficile, 906 Toxin neutralization assays, Corynebacterium diphtheriae, 496 Toxin tests, for Clostridium, 956 Toxins Bacillus anthracis, 445, 455 Clostridium botulinum, 946–947, 952 Clostridium difficile, 944, 951 Clostridium histotoxic skin and soft tissue infections, 944–946 Clostridium perfringens enterotoxin, 942– 943 Clostridium perfringens major extracellular, 945–946 Clostridium septicum, 946 Clostridium sordellii, 946 Clostridium tetani, 947–948, 952 community-acquired respiratory distress syndrome (CARDS), 1088–1089, 1094 diphtheria, 480, 488, 496 enterotoxin Bacteroides fragilis, 970 Clostridium perfringens, 942–943 enterotoxigenic Escherichia coli (ETEC), 685–686, 688–690, 692–693, 695–697 Staphylococcus, 360, 366 Escherichia coli enterotoxigenic (ETEC), 685–686, 688– 690, 692–693, 695–697 Shiga toxin-producing (STEC), 685, 688–692, 694, 696–697 Hafnia, 721 mycotoxins, 2188–2192 aflatoxins, 2188–2189 bioterrorism, 2192 chemical classification and biosynthesis, 2188 citrinin, 2189 cyclopiazonic acid, 2189 ergot alkaloids, 2189–2190 food safety, 2190–2192 fumonisins, 2189–2190 ochratoxins, 2189–2190 patulin, 2189–2190 sick building syndrome, 2192 taxonomy of mycotoxin-producing fungi, 2191 trichothecenes, 2189–2190 zearalenone, 2189–2190 Shigella, 698 T-2 toxin, 2190 Vibrio cholerae, 763, 768 Toxocara, 2448, 2495–2496 clinical significance, 2496 description of agents, 2495 direct examination by microscopy, 2494, 2496 epidemiology, transmission, and prevention, 2495–2496

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 149

serologic tests, 2496 treatment, 2496, 2531 Toxocara canis, 2495 commercial kits for immunodetection of serum antibodies, 2296 detection, 2328, 2330 Toxocara cati, 2328, 2330, 2495 Toxocariasis, 2330, 2495–2496 commercial kits for immunodetection of serum antibodies, 2296 treatment, 2532 Toxoplasma gondii, 2373–2382 antigen detection, 2375 antimicrobial susceptibilities, 1183 cell-mediated immune responses, 2376 clinical significance, 2375 clinical use of immunodiagnostic tests, 2378–2381 determination of immune status, 2378– 2379 diagnosis during pregnancy, 2379–2380 diagnosis in immunocompromised hosts, 2381 diagnosis in newborns, 2380–2381 diagnosis of acute acquired infections, 2379 diagnosis of ocular infection, 2381 collection, transport, and storage of specimens, 2375 collection for antibody determination, 2375 collection for determination of parasite DNA, 2375 commercial kits for immunodetection of serum antibodies, 2296 culture, 2307 detection, 2327–2331 direct examination, 2375–2376 epidemiology and transmission, 2373–2375 isolation procedures, 2376 life cycle, 2373–2374 microscopy, 2375 nucleic acid detection techniques, 2375– 2376 prevention, 2374–2375 serologic tests, 2376–2379 taxonomy, 2373 treatment, 2381–2382, 2530 Toxoplasmatinae (subfamily), 2373 Toxoplasmosis, 2373–2382 commercial kits for immunodetection of serum antibodies, 2296 congenital, 2373, 2375, 2379–2380 detection, 2331 ocular, 2375, 2381 TORCH (toxoplasmosis, other, rubella, cytomegalovirus, and herpes simplex virus) panels, 1530 TPHA test, 1066, 1070 TP-PA test, 1066, 1070, 1074 Trabulsiella guamensis, 718, 727 Tracheal aspirate, Gram stain and plating medium recommendations, 286 Tracheitis, etiologies of, 290 Tracheobronchitis Mycoplasma, 1091 respiratory syncytial virus (RSV), 1500 Trachipleistophora, 2209–2211, 2328–2330 Trachipleistophora anthropophthera, 2210, 2213, 2328, 2330 Trachipleistophora hominis, 2209–2210, 2213, 2215, 2329, 2332 Trachoma, 2505, 2513 Trans-acting transcriptional activator (Tat) protein/gene, HIV, 1436–1437

n cxlix

Transcription-mediated amplification (TMA), 63–64 hepatitis C virus, 1603–1604 human papillomavirus (HPV), 1793 Trichomonas vaginalis, 2415 Transduction, 1216 Transformation, 1216 Transfusion-related infections, Yersinia enterocolitica and, 742 Transient aplastic crisis, 1818–1819, 1822 Transmissible spongiform encephalopathies (TSEs), 1859–1864 antigen detection, 1863–1864 clinical significance, 1861 collection, transport, and storage of specimens, 1861–1862 safety and security, 1861–1862 shipping, 1862 specimen collection, 1862 CSF analysis, 1864 description of agent, 1859 detection and identification methods, 1435 direct examination, 1862–1864 epidemiology and transmission, 1859–1861 evaluation, interpretation, and reporting of results, 1864 microscopy, 1862–1863 mutations in PRNP, 1862 nucleic acid detection, 1863–1864 taxonomy, 1859 Transmission, see specific organisms Transmission electron microscopy, see Electron microscopy Transplantation patients Acanthamoeba, 2389, 2392 adenoviruses, 1771–1773, 1775, 1777 Aspergillus, 2033 Blastoschizomyces, 1992 Cystoisospora belli, 2428 cytomegalovirus, 1719 Epstein-Barr virus, 1739–1740, 1746–1747 Fusarium, 2065, 2067 hepatitis B virus, 1844 human herpesvirus 6 (HHV-6), 1756, 1760 human herpesvirus 7 (HHV-7), 1761 human herpesvirus 8 (HHV-8), 1763 Leishmania, 2359 mucormycosis, 2088–2089 polyomaviruses, 1804–1805, 1812 respiratory syncytial virus, 1500–1501 Trypanosoma cruzi, 2363 Transport medium, Stuart, 345 Transport of samples, 178; see also Specimen collection, transport, and processing anaerobic bacteria, 905 Mycobacterium, 546–547 parasitology, 2311 blood, 2304 stool, 2293–2294, 2296 specimen maintenance during, 282 viruses, 1405–1417 media for viral cultures, 1429–1430 transport conditions, 1410 transport medium, 1409–1410 Transposons, 1217, 1220, 1229–1230, 1234, 1325 Transverse myelitis, Mycoplasma and, 1091 Trapdoor spider, 2520 Trauma-related infection Actinobacillus, 654 Mycobacterium, 596, 598 Mycobacterium marinum, 542

cl

n

SUBJECT INDEX

Trauma-related infection (continued) Mycobacterium ulcerans, 543 Nocardia, 515–516 Nocardia brasiliensis, 516 Pantoea, 719 Stenotrophomonas maltophilia, 794 Vibrio vulnificus, 765 Trehalose, 2001 TREK Sensititre MYCOTB MIC plate method, 1367 Trematoda (class), 2290 Trematodes (flukes), 2479–2490 detection, 2323 foodborne digeneans, 2484, 2487–2490 intestinal, 2482, 2484, 2490 life cycles, 2479, 2483–2484 liver, 2481, 2484, 2487–2490 of circulatory system, 2479–2486; see also Schistosoma anthelminthic susceptibility and treatment, 2486 clinical significance, 2484–2485 collection, transport, and storage, 2486 detection, 2486 epidemiology and transmission, 2484 serology, 2486 taxonomy, 2479–2480, 2484 typing, 2486 respiratory system, 2481, 2484, 2487 taxonomy and classification, 2288, 2290 treatment, 2531, 2533 Trematosphaeria grisea, 1965, 1968, 2174, 2176, 2178–2180, 2182 Trematosphaeria pertusa, 2174 Trematosphaeriaceae (family), 2174 Tremellomycetes (class), 1937 Trench fever, 876, 2507, 2511 Trepanostika Tp Recombinant, 1068 Trep-Chek IgG, 1067 Trep-Chek IgM, 1067, 1071 Trepomonadea (class), 2287, 2408–2409 Treponema, 1055–1075 antimicrobial susceptibilities, 1072 clinical significance, 1059–1061 endemic treponematoses, 1061 venereal syphilis, 1059–1061 collection, transport, and storage of specimens, 1061–1063 oral treponemes (T. denticola), 1063 syphilis (T. pallidum), 1061–1063 description of agents, 1055–1057 direct examination, 1063 endemic treponematoses, 1055–1056, 1058, 1061 epidemiology and transmission, 1057–1059 endemic treponematoses, 1058 oral treponemes, 1058–1059 venereal syphilis, 1057–1058 evaluation, interpretation, and reporting of results, 1072–1075 direct detection of Treponema pallidum, 1072–1073 serologic tests, 1073–1074 syphilis tests in HIV infection, 1075 tests for congenital syphilis, 1075 tests for neurosyphilis, 1074 identification, 994, 997 isolation procedures, 1064 oral treponemes collection, transport, and storage of specimens, 1063 description, 1057 epidemiology and transmission, 1058– 1059

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 150

serologic tests, 1065–1072 taxonomy, 1055 treponemal tests for syphilis, 1066–1072 chemiluminescence immunoassays (CLIAs), 1071–1072 combined treponemal IgM/IgG EIAs, 1068 conventional, 1066, 1070, 1074 EIAs, 1067–1068, 1070–1071, 1074 FTA-ABS test, 1066, 1070, 1074 immunoblot assays, 1070, 1072 MHA-TP test, 1066, 1070, 1074 multiplex flow immunoassays, 1071– 1072 rapid combination nontreponemal and treponemal tests, 1070 rapid treponemal tests, 1069, 1071– 1072, 1074 TPHA, 1066, 1070 TP-PA test, 1066, 1070, 1074 treponemal IgM or IgG EIAs, 1067 typing systems, 1065 Treponema amylovorum, 1057 Treponema carateum, 1055–1056 Treponema denticola, 229, 1055, 1057–1059 Treponema lecithinolyticum, 1057 Treponema maltophilum, 1057 Treponema medium, 1057 Treponema pallidum, 997 antimicrobial susceptibilities, 1072, 1187 clinical significance, 1059–1061 collection, transport, and storage of specimens, 1061–1063 dark-field microscopy, 296–297, 1055, 1062–1063, 1072 description, 1055, 1057 direct detection, 1072–1073 direct examination, 1063 epidemiology and transmission, 1057–1058 evaluation, interpretation, and reporting of results, 1072–1075 isolation procedures, 1064 morphology, 1057 subspecies, 1055 treponemal tests for syphilis, 1066–1072 chemiluminescence immunoassays (CLIAs), 1071–1072 combined treponemal IgM/IgG EIAs, 1068 conventional, 1066, 1070, 1074 EIAs, 1067–1068, 1070–1071, 1074 FTA-ABS test, 1066, 1070, 1074 immunoblot assays, 1070, 1072 MHA-TP test, 1066, 1070, 1074 multiplex flow immunoassays, 1071– 1072 rapid combination nontreponemal and treponemal tests, 1070 rapid treponemal tests, 1069, 1071– 1072, 1074 TPHA, 1066, 1070 TP-PA test, 1066, 1070, 1074 treponemal IgM or IgG EIAs, 1067 typing systems, 1065 Treponema pallidum μ-capture IgM ELISA, 1067 Treponema pallidum subsp. endemicum, 1055– 1056 Treponema pallidum subsp. pallidum, 1055– 1056 Treponema pallidum subsp. pertenue, 1055– 1056 Treponema parvum, 1057 Treponema pectinovorum, 1057

Treponema pertenue, 1055 Treponema putidum, 1057 Treponema socranskii, 1057 Treponema vincentii, 1057 Treponema+VDRL ViraBlot, 1070 Trep-Sure, 1068 Tretinoin, 2515 Triage parasite panel, 2295 Triatoma, 2507 Triatoma rubrofasciata, 2508 Triatoma sanguisuga, 2508 Trichinella, 2495 clinical significance, 2495 description, 2495 detection, 2329, 2332 direct examination by microscopy, 2494– 2495 epidemiology, transmission, and prevention, 2495 nucleic acid detection, 2495 serologic tests, 2495 treatment, 2495 Trichinella britovi, 2495 Trichinella pseudospiralis, 2495 Trichinella spiralis, 2495 commercial kits for immunodetection of serum antibodies, 2296 life cycle, 2495 microscopy, 2494 nucleic acid detection, 2495 treatment, 2531 Trichinellidae (family), 2289 Trichinelloidea (superfamily), 2289, 2495 Trichinosis (trichinellosis), 2296, 2332, 2495 Trichobilharzia, 2480 Trichocomaceae (family), 2073 Trichodectes canis, 2510 Trichoderma, 2058, 2073, 2076 key phenotypic features, 2065 trichothecenes, 2190 Trichoderma citrinoviride, 2065, 2073 Trichoderma harzianum, 2065 Trichoderma longibrachiatum, 2065, 2073, 2075, 2271 Trichoderma pseudokoningii, 2073 Trichodysplasia spinosa, 1805–1807 Trichodysplasia spinosa-associated polyomavirus (TSPyV), 1803, 1805– 1806, 1810 TrichOKEY, 2073 Trichomaceae (family), 2030 Trichomonadea (class), 2287, 2408, 2412– 2413 Trichomonadida (order), 2287, 2412–2413 Trichomonas Aptima Combo 2 assay, 2415 Trichomonas culture system, 2315, 2327 Trichomonas hominis, 2400, 2408–2409, 2416 Trichomonas tenax, 2400 detection, 2329, 2331 sputum specimen, 2305 Trichomonas vaginalis, 2399–2400, 2413–2416 antigen detection, 2327, 2415 antiparasitic agent resistance, 2553–2554 antiparasitic agent susceptibility testing methods, 2564, 2566 clinical significance, 2414 culture, 2307, 2414–2415 description, 2413 detection, 2326–2327 direct detection, 2414–2415 epidemiology, transmission, and prevention, 2413–2414 evaluation, interpretation, and reporting of results, 2415–2416

SUBJECT INDEX media for culture, 2315–2316 metronidazole resistance, 1232 microscopy, 2414 nucleic acid detection, 2308, 2415 specimens, 2305 taxonomy, 2413 treatment, 1194, 2415, 2530, 2535, 2544– 2545 Trichomoniasis, 2414–2416 antiparasitic agent resistance mechanisms, 2553–2554 antiparasitic agent susceptibility testing methods, 2566 Trichomonascus ciferrii, 1985, 2000 Trichophyton, 1937, 1939, 2128–2146 anatomic specificity, 2136 anthropophilic species, 2135 antifungal susceptibility testing, 2271 antimicrobial susceptibilities, 2145 characteristics, 2130–2134 clinical significance, 2135–2136 colony characteristics, 2140, 2144 description of etiologic agents, 2145 epidemiology and transmission, 2135 evaluation, interpretation, and reporting of laboratory results, 2146 geophilic species, 2135 growth on BCPMSG, 2141 identification, 2139–2141 in vitro hair perforation test, 2140 isolation, 2138–2139 laboratory testing of specimens, 2137– 2139 media, 1959–1962 microscopy, 1975, 2137–2142 molecular identification techniques, 2141 nucleic acid detection, 2139 nutritional requirements, 2140 physiological tests, 2140–2141 specimen collection, transport, and processing, 1944, 1947, 1953, 2136–2137 strain typing systems, 2145 taxonomy, 2128 temperature tolerance and temperature enhancement, 2141 urea hydrolysis, 2141 zoophilic species, 2135 Trichophyton agars 1 to 7, 1962 Trichophyton ajelloi, 2130, 2141, 2145 Trichophyton bullosum, 2131 Trichophyton concentricum, 2131, 2135, 2140 Trichophyton eboreum, 2133 Trichophyton equinum, 2128, 2131, 2135 Trichophyton equinum var. autotrophicum, 2140 Trichophyton equinum var. equinum, 2140 Trichophyton erinacei, 2131, 2135 Trichophyton eriotrephon, 2131 Trichophyton fluviomuniense, 2132 Trichophyton interdigitale, 2131–2132, 2136, 2144–2145 Trichophyton kanei, 2132, 2145 Trichophyton krajdenii, 2132, 2144 Trichophyton megninii, 2131, 2135, 2138, 2140 Trichophyton mentagrophytes complex, 1960, 2131–2132, 2135–2136, 2138, 2140– 2142, 2144–2145, 2262 Trichophyton raubitschekii, 2132, 2136, 2141– 2142 Trichophyton rubrum, 2118, 2132, 2135, 2139–2142, 2144–2146 antifungal susceptibility testing, 2262, 2268–2269, 2271

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 151

media, 1960 Trichophyton schoenleinii, 2133, 2135–2136, 2138–2139, 2145 Trichophyton simii, 2133, 2135 Trichophyton soudanense, 1947, 2133, 2135, 2137–2138, 2140–2141, 2145 Trichophyton terrestre, 2133, 2139, 2141– 2142, 2146 Trichophyton thuringiense, 2133 Trichophyton tonsurans, 2128, 2133, 2135– 2138, 2140, 2142, 2144–2145 Trichophyton vanbreuseghemii, 2133, 2135 Trichophyton verrucosum, 2134–2136, 2138, 2140, 2142, 2145 Trichophyton violaceum, 2134–2136, 2138, 2140 Trichophyton yaoundei, 2140 Trichosporon, 1937, 1986, 1997–1998, 2004 antifungal susceptibilities, 2005–2006 antifungal susceptibility testing, 2263 characteristics of, 1991 clinical significance, 1994 cycloheximide inhibition, 1955 description of agents, 1991 identification, 2000, 2003 media, 1960 microscopic appearance on morphology agar, 1998 morphologic features, 1987 specimen collection, transport, and processing, 1947–1949 taxonomy, 1985 white piedra, 2148 Trichosporon asahii, 1951, 1985, 1989, 1991, 1994, 2005, 2148 Trichosporon asteroides, 1985, 1991, 1994, 2005 Trichosporon beigelii, 1985, 2148 Trichosporon capitatum, 1984 Trichosporon cutaneum, 1985, 1991, 1994, 2005, 2148 Trichosporon dermatis, 1985 Trichosporon inkin, 1985, 1991, 1994, 2148 Trichosporon japonicum, 1994 Trichosporon jirovecii, 1985 Trichosporon loubieri, 1985, 1991, 1994 Trichosporon mucoides, 1985, 1989, 1991, 1994, 2005 Trichosporon mycotoxinivorans, 1985, 1991, 1994 Trichosporon ovoides, 1985, 1989, 1991, 1994, 2005, 2148 Trichosporonosis, 1947, 1994 Trichostomatia (order), 2287, 2416 Trichostrongylidae (family), 2289 Trichostrongyloidea (superfamily), 2289 Trichostrongylus detection, 2320, 2322–2323 eggs, 2449 treatment, 2531, 2535 Trichothecenes, 2189–2192 Trichothecium, 2190 Trichrome stain, 2314 for parasites, 2318–2319 modified, 2319 Trichuridae (family), 2289, 2497 Trichuris trichiura, 2458–2459 clinical significance, 2458–2459 description, 2459 eggs, 2449, 2451, 2459 larvae, 2459 worms, 2459 detection, 2320, 2323 diagnosis, 2458

n cli

epidemiology and prevention, 2459 taxonomy, 2459 transmission and life cycle, 2454, 2459 treatment, 2455, 2459, 2531–2532, 2534– 2535 Triclabendazole Fasciola, 2490 Paragonimus, 2487 Triclosan, 186 Tri-Combo parasite screen, 2411 Trifluridine herpes simplex virus (HSV), 1689 herpesviruses, 1884, 1886 spectrum of activity, 1885 Trimethoprim, 1191–1192 adverse effects, 1192 antimicrobial susceptibility testing, 1256, 1261 concentration in serum, 1199 mechanism of action, 1191 pharmacology, 1191 spectrum of activity, 1191–1192 with sulfonamides, 1191–1192, 1199 Trimethoprim resistance, 1234, 1385 Trimethoprim-sulfamethoxazole antimicrobial susceptibility testing, 1256, 1261.1266 Blastocystis hominis, 2407 Cyclospora cayetanensis, 2431 Cystoisospora belli, 2430–2431 Pneumocystis, 2025–2026 Toxoplasma gondii, 2382 Trimethoprim-sulfamethoxazole resistance, 1234–1235 acquired resistance to sulfonamides, 1234– 1235 acquired resistance to trimethoprim, 1234 common associations of resistance mechanisms, 1215 intrinsic resistance, 1234 Streptococcus pneumoniae, 1319 Triple-centrifugation method for trypanosomes, 2336 tRNAscan-SE, 233 Trofile coreceptor tropism, 1448 Troglotrematidae (family), 2290, 2481, 2482, 2487, 2490 Trombidiformes (order), 2511 Tropheryma whipplei, 75, 240, 1159–1165 antimicrobial susceptibilities, 1163–1164 clinical significance, 1083, 1160–1161 classical (systemic) Whipple’s disease, 1160 isolated organ manifestations, 1160– 1161 pathogenesis and immunology, 1160 symptoms and signs, 1160 collection, transport, and storage of specimens, 1161–1162 culture, 1159 description of, 1159 diagnostic tests, 1087 direct examination, 1162–1163 antigen detection, 1162 microscopy, 1162 nucleic acid detection, 1162–1163 epidemiology and transmission, 1083, 1159 evaluation, interpretation, and reporting of results, 1164–1165 genome, 1159 identification, 1163 isolation procedures, 1163 morphology, 1159

clii

n

SUBJECT INDEX

Tropheryma whipplei (continued) PAS staining, 1159–1160 serologic tests, 1087, 1163 taxonomy, 1159 Tropical pulmonary eosinophilia, 2464 Trovafloxacin, 1179, 2382 Trovagene HPV test, 1788, 1790 TRU FLU, 1474 Trueperella, 474 clinical significance, 479 description of genus, 478 epidemiology and transmission, 478–479 identification, 438, 496 isolation procedures, 480 taxonomy, 474–475 Trueperella bernardiae, 478–479, 484, 496 Trueperella pyogenes, 478–479, 484, 496 Trugene HBV, 1920 Trugene HCV, 1606 Trugene HIV-1, 1447–1448, 1450 TruGene HIV-1 genotyping assay, 1920 Trypanosoma, 2362–2368 blood specimen, 2307 culture, 2307 detection, 2333 media for culture, 2315–2316 stains for detection, 2312–2313 storage methods, 166 Trypanosoma brucei, 2366–2368 animal inoculation, 2367 antiparasitic agent resistance, 2555 antiparasitic agent susceptibility testing methods, 2564, 2567 arthropod vector, 2507 clinical significance, 2366 collection of specimens, 2367 culture, 2367 diagnosis, 2367 direct examination, 2367 epidemiology and transmission, 2366 life cycle and morphology, 2366 microscopic detection, 2367 PCR detection, 2367 prevention, 2367–2368 serologic tests, 2367 treatment, 2367–2368, 2530, 2542–2545 Trypanosoma brucei gambiense, 2364, 2366– 2368 antiparasitic agent resistance, 2555 characteristics of, 2360 detection, 2328, 2367 taxonomy, 2357 treatment, 2367–2368, 2543–2545 Trypanosoma brucei rhodesiense, 2366–2368 antiparasitic agent resistance, 2555 characteristics of, 2360 detection, 2328, 2367 taxonomy, 2357 treatment, 2367, 2543–2545 Trypanosoma congolense, 2368 Trypanosoma cruzi, 2362–2365 animal inoculation, 2365 arthropod vector, 2507–2508 bone marrow aspirate, 2306 characteristics of, 2360 clinical significance, 2362–2364 collection of specimens, 2364 commercial kits for immunodetection of serum antibodies, 2296 culture, 2365 detection, 2327–2329 diagnosis, 2364 direct examination, 2364–2365 epidemiology and transmission, 2362

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 152

life cycle and morphology, 2362–2363 media for culture, 2315 microscopic detection, 2364–2365 PCR detection, 2365 prevention, 2365 serologic tests, 2365 taxonomy, 2357 treatment, 2365, 2530, 2542, 2545 xenodiagnosis, 2307, 2365 Trypanosoma evansi, 2357, 2368 Trypanosoma lewisi, 2368 Trypanosoma rangeli, 2360, 2365, 2508 Trypanosomatida (order), 2287 Trypanosomes antiparasitic agent resistance mechanisms, 2555 arthropod vectors, 2508 detection, 2334–2336 triple-centrifugation method, 2336 Trypanosomiasis antiparasitic agent susceptibility testing methods, 2564, 2567 characteristics of, 2360 detection, 2331 Trypanozoon (subgenus), 2357 Trypsin activity, 616 Trypsin solutions, 1423 Tryptic soy blood agar, 345 Trypticase soy agar, with sheep blood, sucrose, and tetracycline, 346 Trypticase soy agar, with sheep blood and gentamicin, 346 Trypticase soy agar, with sheep blood and vancomycin, 346 Trypticase soy agar with 5% sheep, rabbit, or horse blood, 2315 Trypticase soy broth, 17, 345–346 Trypticase tellurite agar base, 346 TSEs, see Transmissible spongiform encephalopathies Tsetse flies, 2505–2506 TSO3 OZO-TEST, 205 T-Spot.TB assay, 555–556, 576 TSST-1 Evigene, 366 TST-RPLA, 366 Tsukamurella acid-fast stain, 321 chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 519 collection, transport, and storage, 520 description of genus, 511 G+C content, 536 identification, 438, 522, 524, 526–527 microscopy, 521 morphologic characteristics, 508 taxonomy, 504–505 Tsukamurella inchonensis, 514 Tsukamurella paurometabola, 511, 513, 519 Tsukamurella pulmonis, 510, 512, 514, 527 Tsukamurella spumae, 514, 527 Tsukamurella strandjordii, 510, 514 Tsukamurella tyrosinosolvens, 513–514, 519, 527 T-Track CMV ELISpot, 1730 Tubal factor infertility, Chlamydia trachomatis and, 1108 Tuberculin skin test, 546, 555–556, 576 Tuberculosis, 536, 538 antimicrobial susceptibilities, 1180, 1356– 1368 antimicrobial susceptibility testing, 1361– 1368

microscopic observation of drug susceptibility (MODS), 1356, 1367 molecular methods, 1356, 1367–1368 BCG vaccine, 538, 555 drug-resistant, 1356, 1360–1364, 1367– 1368 extensively drug-resistant (XDR-TB), 536, 538, 583, 1356, 1367–1368 HIV-associated, 576 interferon gamma release assays (IGRAs), 546, 555–556, 576 laboratory-acquired infections, 177 multidrug-resistant (MDR-TB), 536, 576, 583–584, 1356, 1360–1362, 1367 treatment, 1356–1368 antimicrobial agents, 1356–1361 antimicrobial susceptibility testing, 1361–1368 DOTS (directly observed therapy, short-course), 536 tuberculin skin test (TST), 546, 555–556, 576 Tubex test, 704 Tubo-ovarian abscess, Neisseria gonorrhoeae and, 636 Tubulinosema, 2209–2210 Tubulinosema acridophagus, 2210, 2212 tuf gen, Enterococcus, 405, 411 Tularemia, 222, 851, 854–855, 2507; see also Francisella tularensis Tumbu fly, 2517 Tunga penetrans, 2509–2510 Tungiasis, 2509–2510, 2516 Turicella antimicrobial susceptibility testing, 1328 description of genus, 475 identification, 438, 494 taxonomy, 474–475 Turicella otitidis, 475, 476, 478, 484–485, 494, 1328 Turicibacter, 921 Turicibacter sanguinis, 922, 926 Turkey X disease, 2188 Turneriella, 1028 Turneriella parva, 1030 Tween 20-PBS, 1423 23S rRNA gene sequence Arcobacter, 1005 Campylobacter, 1005, 1007 Chlamydiaceae, 1106, 1112 Helicobacter, 1023 Legionella pneumophila, 892 nontuberculous mycobacteria (NTM), slowly growing, 580 Treponema, 1072 Twinrix, 1585, 1590 TYI-S-33 medium, 2315 Tymovirales (order), 1402 Typeability, definition, 132 Typhidot test, 704 Typhoid fever, 92, 701–702 Typing, see Subtyping; specific organisms TYSGM-9 medium, 2315 Tzanck test, 1707 U9B broth, 346 UAB Diagnostic Mycoplasma Laboratory, 1094 UL97 gene, cytomegalovirus, 1895–1896 Ulcerative colitis Faecalibacterium prausnitzii, 925 non-spore-forming, anaerobic, Grampositive rods, 923

SUBJECT INDEX Ulcers anaerobic Gram-negative rods, 972 Anaerococcus, 911 Blastomyces dermatitidis, 2114 etiologies, usual, 290 Finegoldia magna, 911 Fusarium, 2058 Gram-positive anaerobic cocci (GPAC), 910–911 herpes simplex virus (HSV), 1688 Histoplasma capsulatum, 2114 parasitology, 2294, 2298 Peptoniphilus, 911 Porphyromonas, 971 Treponema, 1061 Treponema pallidum, 1058 Ultralow-temperature freezing, 162–164 cryoprotective agents, 163–164 freezing method, 164 preparation of microbes for freezing, 164 specialized storage systems, 164 storage vials, 163 thawing, 164 Ultrasound, for lymphatic filarial nematodes, 2464 Uncinocarpus, 2109 Uncultivable organisms anaerobic Gram-negative rods, 983 discovery by molecular methods, 75 in tissues, sample handling, 292 Unheated serum reagin (USR), 1066 Uni-Gold Recombigen HIV-1/2, 1445 Unique recombinant forms (URF), HIV, 1436 United States Preventive Services Task Force (USPSTF), cervical cancer screening recommendations, 1786 University of Orsay, 150 UPEC, see Escherichia coli, uropathogenic Upper respiratory tract infection adenoviruses, 1771–1772 bocavirus, 1823 coronaviruses, 1569 Corynebacterium diphtheriae, 480 human metapneumovirus, 1509 influenza virus, 1471 parainfluenza virus, 1488 respiratory syncytial virus (RSV), 1500 rhinoviruses, 1553 Upper respiratory tract specimen collection, transport, and handling, 279, 299–301 A. haemolyticum, 300 C. diphtheriae, 300–301 epiglottitis, 301 nasal, 300 nasopharynx, 300 sinus, 300 throat, 299–300 fungi, 1946–1948, 1950 viruses, 1408 Uracil-N-glycosylase (UNG), 79 ureA, Helicobacter, 1019, 1021, 1023 Urea agar, 346 Urea breath test, 1019, 1023 Urea hydrolysis dermatophytes, 2141 Streptococcus, 395 Ureaplasma, 1088–1101 A7 agar for, 325 antimicrobial susceptibilities, 1099–1100 biosafety considerations, 1095 clinical significance, 1083, 1091–1093 collection, transport, and storage of specimens, 1093

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 153

colonies, 1090 description of, 1088–1090 diagnostic tests, 1087 direct examination, 1094–1095 epidemiology and transmission, 1083, 1090–1091 evaluation, interpretation, and reporting of results, 1100–1101 identification, 1096 isolation procedures, 1095–1096 serologic tests, 1087, 1098 taxonomy, 1089 typing systems, 1096–1097 Ureaplasma parvum clinical significance, 1091–1092 evaluation, interpretation, and reporting of results, 1101 nucleic acid detection, 1094–1095 Ureaplasma urealyticum A7 agar for, 325 A8 agar, 325, 327 A8 agar for, 327 antimicrobial susceptibilities, 1180, 1183, 1186–1187 clinical significance, 1083, 1091–1093 diagnostic tests, 1087 epidemiology and clinical diseases associated with, 1083 evaluation, interpretation, and reporting of results, 1100–1101 nucleic acid detection, 1094–1095 urogenital Mycoplasma broth base for, 346 Urease test, 319 yeasts, 2000–2001 Yersinia enterocolitica, 742 Ureidopenicillins, 1173 Ureteral stenosis, BK polyomavirus and, 1804 Urethra specimens, 278 Urethral secretion specimens, 277 Urethral syndrome, etiologies of, 291 Urethritis adenoviruses, 1773 Chlamydia trachomatis, 1108 etiologies, usual, 290 Mycoplasma genitalium, 1101 Neisseria gonorrhoeae, 636 Neisseria meningitidis, 637 Trichomonas vaginalis, 2414 viruses, specimens and methods for detection of, 1408 Urinary tract infection (UTI) Actinobaculum, 924 Actinomyces schaalii, 924 Aerococcus urinae, 431 Aeromonas, 754 Alloscardovia omnicolens, 925 anaerobic Gram-negative rods, 972 Arthrobacter, 479 Atopobium vaginae, 925 Bacillus cereus, 443 Campylobacter, 1000 Candida, 1993 Citrobacter, 720 Corynebacterium amycolatum, 479 Corynebacterium aurimucosum, 479 Corynebacterium minutissimum, 479 Corynebacterium riegelii, 479, 492 Corynebacterium urealyticum, 479, 493 Enterococcus, 406, 415 Escherichia coli, 686 etiologies, usual, 291 fungal, 1950 Globicatella, 425

n cliii

Gram-positive anaerobic cocci (GPAC), 910–911 Lactococcus, 424 microsporidia, 2210 Mycoplasma, 1092 Myroides odoratus, 824 non-spore-forming, anaerobic, Grampositive rods, 923 Oligella, 821 Pasteurella, 655 Proteus, 720 Pseudomonas aeruginosa, 775 Salmonella, 701 Serratia, 720 Staphylococcus, 360 Stenotrophomonas maltophilia, 794 Urinary tract infection chromogenic agar (UTIC agar), 346 Urinary tract specimen collection, 271 Urine culture results interpretation, 305 specimen handling, 303–304 Urine specimen collection of specimens, 281 fungi, 1946–1948, 1951 Gram stain and plating medium recommendations, 286 Mycobacterium, 548 sample collection, transport, and storage guidelines, 279–280, 303–305 screening specimens, 284 viruses, 1416 Urogenital Mycoplasma broth base, 346 Urogenital specimen antigen detection, 2327 culture, 2327 direct wet mount, 2326–2327 parasitology, 2294, 2300, 2305, 2326–2327 Urogenital tract infections anaerobic Gram-negative rods, 972 viruses, specimens and methods for detection of, 1408 Uroplectes, 2520 Urosepsis, Aerococcus and, 424 Urticaria carbapenems, 1177 polymyxins, 1193 quinolones, 1180 rifaximin, 1195 tetracyclines, 1187 Urticating hairs caterpillars, 2518–2519 spider, 2521 U.S. Army Medical Research Institute of Infectious Diseases (USAMRIID), 218 U.S. Army Medical Research Unit (USAMRU) medium, 2315 U.S. Department of Agriculture (USDA) Agricultural Select Agent Program, 219 Animal and Plant Health Inspection Service (APHIS), 219 outbreak response, 121 U.S. Department of Health and Human Services (HHS) biothreat agents and, 217–218 outbreak response, 121 U.S. Department of Transportation, 1416 USA Patriot Act, 219 Ushijima’s medium, 1953 Ustilaginaceae (family), 1985, 2075 Ustilaginomycotina (subphylum), 1985 Ustilago maydis, 1985

cliv

n

SUBJECT INDEX

Uveitis human T-cell lymphotropic viruses (HTLVs), 1461 Mycobacterium kansasii, 542 Mycobacterium simiae, 543 Onchocerca volvulus, 2466 viruses, specimens and methods for detection of, 1407 UVM (University of Vermont) modified Listeria enrichment broth, 346 V agar, 346 V factor, 667, 672–673, 675 V8 agar, 1962 VACC agar (Remel VACC agar), 346 Vaccine adenoviruses, 1772 arboviruses, 1645–1647 Bacillus anthracis, 447 Bacillus Calmette-Guérin (BCG), 538, 555, 576 Bordetella pertussis, 845 Borrelia, 1048 Brucella, 869 Clostridium tetani, 948 controversy, 1521 cytomegalovirus, 1720 Giardia, 2411 Haemophilus influenzae type b (Hib), 668– 669, 677 hantavirus, 1662 hepatitis A virus, 1584–1585, 1589–1590 hepatitis B virus, 1585, 1843, 1845, 1849– 1851, 1853 hepatitis E virus, 1586, 1590 human immunodeficiency virus, 1440, 1449 human papillomavirus (HPV), 1786 human T-cell lymphotropic viruses (HTLVs), 1461 influenza viruses, 1471–1472 measles, 1519–1521 MMR (measles-mumps-rubella), 1492– 1493, 1521 mumps virus, 1492–1493 parainfluenza virus, 1488 parvovirus B19, 1818 poliovirus, 148, 1539–1540, 1544–1545, 1804 protection against laboratory-acquired infectious diseases, 176 Q fever (QVAX), 1155 rabies, 1635–1636 rotaviruses, 1622 rubella virus, 1526 Schistosoma, 2486 secondary vaccine failure, 1520 smallpox, 1829, 1831 vaccinia, 1829, 1831 varicella-zoster virus, 1704, 1706, 1709 Vaccine Preventable Disease Reference Centers (VPD RCs), 1523 Vaccine reactions, molecular methods to identify, 147–148 Vaccinia necrosum, 1831 Vaccinia virus cell culture, 1422 clinical significance, 1831 cytopathic effect (CPE), 1836 epidemiology and transmission, 1828–1829 PCR assay, 1834 serologic tests, 1836–1837 vaccine, 1829, 1831

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 154

Vacuolating cytotoxin (vacA) gene, Helicobacter, 1021 Vagabond’s disease, 2510–2511 Vaginal candidiasis, 1951 Vaginal microbiome, 231 Vaginal specimens fungi, 1946, 1951 parasitology, 2300 specimen collection, transport, and storage guidelines, 277 Vaginitis Candida, 2327 Candida africana, 1992 Gardnerella vaginalis, 2327 specimen collection, transport, and handling, 296 Trichomonas vaginalis, 2327, 2414 Vagococcus antimicrobial susceptibilities, 430 clinical significance, 424 identification, 425–426, 426, 429 taxonomy, 422–423 Vagococcus fluvialis, 409–410, 422, 424 Vahlkampfia, 2387 Vahlkampfiidae (family), 2387 Valacyclovir cytomegalovirus, 1720 Epstein-Barr virus, 1740 herpes B virus, 1697 herpes simplex virus (HSV), 1689, 1919 herpesviruses, 1882–1883, 1886 varicella-zoster virus, 1706 Valacyclovir resistance, 1895, 1917 Valganciclovir cytomegalovirus, 1720 Epstein-Barr virus, 1740 herpesviruses, 1884–1885 human herpesvirus 8 (HHV-8), 1763 multicentric Castleman’s disease, 1763 Valganciclovir resistance, 1917 Validation of methods, 149 of molecular methods, 79 Valley fever, 2114 Valovirus (genus), 1617 van genes, 303, 407, 409, 412–415, 1229– 1230, 1249, 1278, 1288–1289, 1328, 1381 Van Leeuwenhoek, Antonie, 11, 238 Vancomycin, 1187–1189 antimicrobial susceptibility testing, 1256, 1261 concentration in serum, 1199 Vancomycin agar screening test, 1289 Vancomycin resistance, 1188, 1214, 1229– 1230 Arcanobacterium haemolyticum, 1328 Clostridium, 1348 detection by automated antimicrobial susceptibility testing, 1278 detection Staphylococcus aureus, 1294–1295 BHI-V6 screening agar test method, 1295 limitations of methods, 1295 quality control, 1295 reporting of results, 1295 Erysipelothrix rhusiopathiae, 1328 in enterococci, 1278, 1288–1289 in streptococci, 1320 Lactobacillus, 1348 molecular methods for detecting in Enterococcus, 1381–1382 Oerskovia turbata, 1328 phenotypic methods for detecting

in Enterococcus, 1288–1289 in Staphylococcus, 1290, 1294–1295 vancomycin agar screening test, 1289 Vancomycin susceptibility, for aerobic Gram-negative bacteria identification, 616 Vancomycin-intermediate S. aureus (VISA), 369–370, 1230 antimicrobial susceptibilities, 1175, 1188– 1189 automated system detection of, 1279 detection of, 1266, 1279, 1294–1295 disk diffusion test for, 1266 hVISA (heterogeneous VISA), 1230, 1295 reporting of, 1295 Vancomycin-intermediate Staphylococcus species (VISS), 1294 Vancomycin-resistant enterococci (VRE) antimicrobial susceptibilities, 413–415 collection, transport, and storage of specimens, 407 commercial sources of chromogenic agar media for, 327 detection, 111, 407, 1288–1289 health care-associated, 406–407 isolation procedures, 408–409 media for detection, 324 molecular detection of antibacterial resistance, 1381–1382 rectal swab screening for, 303 reporting to infection prevention program, 112 surveillance cultures, 113–115 Trypticase soy agar, with sheep blood and vancomycin for, 346 Vancomycin-resistant lactobacilli, 924, 931 Vancomycin-resistant S. aureus (VRSA) antimicrobial susceptibilities, 1175, 1189 automated system detection of, 1279 detection of, 1279, 1294–1295 reporting of, 1295 resistance mechanisms, 1230 Vaniprevir resistance, 1902 Vannella, 2387 VanR assay, 1381–1382 Vapendavir, for rhinoviruses, 1558 Vaqta, 1584, 1590 Variable-number-tandem-repeat (VNTR) analysis, 140 Clostridium, 957 Mycobacterium tuberculosis complex, 584– 585 nontuberculous mycobacteria (NTM), rapidly growing, 603–605 nontuberculous mycobacteria (NTM), slowly growing, 585 Variably protease-sensitive prionopathy (VPSPr), 1859, 1861 Variant Creutzfeldt-Jakob disease (vCJD), 197, 206–207, 1859–1864 Varibaculum, 920–921, 928, 930 Varibaculum cambriense, 920, 923, 926, 928, 930 Varicella congenital varicella syndrome, 1704–1706 epidemiology and transmission, 1704 vaccine-modified disease, 1709 Varicella-zoster immunoglobulin, 1706 Varicella-zoster virus (VZV), 1704–1713 antigen detection, 1707 antimicrobial susceptibilities, 1712 antiviral resistance, 1895, 1917 antiviral susceptibility testing, 1916, 1919, 1924

SUBJECT INDEX congenital varicella syndrome, 1704–1706 cytopathic effect (CPE), 1710 description of agent, 1704 detection and identification methods, 1435 DFA and IFA reagents for the detection of, 1425 diagnostic tests, 1706 direct examination, 1707–1710 electron microscopy, 1705, 1707 epidemiology and transmission, 1704 evaluation, interpretation, and reporting of results, 1712–1713 genome, 1704–1705 genotypes, 1704 identification, 1710–1711 from cell culture, 1710 genetic identification of strains, 1710– 1711 immunofluorescence in H&V-Mix cells, 1429 in immunosuppressed patients, 1705–1706 isolation procedures, 1710 latency, 1704–1705 microscopy, 1707 nucleic acid detection, 1707–1710 PCR, 1706–1710 pregnancy and, 1704–1706 quantitative DNA, 1708 rapid cell culture, 1426 reactivation, 1704–1705, 1709, 1712 serologic tests, 1706, 1711–1712 cellular immunity, 1712 IgG avidity assays, 1711 IgG detection tests in routine diagnostic labs, 1711 IgM tests, 1712 specialized IgG detection tests, 1711 specimen collection and handling, 1406– 1408, 1412, 1415, 1706–1707 taxonomy, 1704 treatment, 1706, 1712 vaccine, 1704, 1706, 1709 Varicellovirus (genus), 1398, 1704 Variola major, 1830 biothreat agent, 221 transmission and disease, 221–222 Variola minor, 1830 Variola virus antiviral therapy, 1831 biothreat agent, 220 characteristics, 220 clinical significance, 1830 epidemiology and transmission, 1828–1829 PCR assay, 1834–1835 serologic tests, 1836 Vascular disease cytomegalovirus, 1719 Pythium insidiosum, 2201 Vasculitis, 2163 Bartonella, 874 BK polyomavirus, 1805 Myceliophthora thermophila, 2076 Sarcocystis, 2431 sulfonamides, 1192 vCJD (variant Creutzfeldt-Jakob disease), 197, 206–207, 1859–1864 VDRL test, 1062, 1066 VDRL-CSF test, 1074–1075 Vectors, arthropod, 2505–2513 Acarina, 2511–2513 Diptera, 2505–2508 Hemiptera, 2508–2509 Phthiraptera, 2510–2511

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 155

Siphonaptera, 2509–2510 VEE (Venezuelan equine encephalitis) virus, 223–224, 1644, 1646–1647 Veillonella, 911 antimicrobial susceptibilities, 916, 1175 antimicrobial susceptibility testing, 1343 clinical significance, 911 description of, 909 epidemiology, 910 identification, 913, 916, 977 isolation procedures, 912 taxonomy, 909 Veillonella alcalescens, 909 Veillonella atypica, 909 Veillonella denticariosi, 909 Veillonella dispar, 909 Veillonella montpellierensis, 977 Veillonella parvula, 909 Veillonella ratti, 977 Veillonella rogosae, 909 Veillonella tobetsuensis, 909 Velvet, 233 Venezuelan equine encephalitis (VEE) virus, 223–224, 1644, 1646–1647 Venezuelan human ehrlichiosis, 1139 Venipuncture, skin disinfection prior to, 17 Venous blood specimen collection, 271 Ventilator-associated pneumonia, Pseudomonas aeruginosa, 774–775 Ventriculitis Bacillus licheniformis, 442 Leuconostoc, 424 Ventriculo-atrial shunt infection, Bacillus subtilis, 442 Venturiales (order), 2153, 2155, 2161, 2163 Verification of molecular methods, 79 Verigene assay, 24, 72 Haemophilus, 677 rhinoviruses, 1555 Verigene Gram-positive blood culture (BCGP) assay, 392 molecular detection of antibiotic resistance, 1381 Staphylococcus, 365 Verigene respiratory virus nucleic acid test (RV+), 1478, 1506 Veronaea, 2153 Verrucarins, 2190, 2192 Verruconis, 2153, 2161, 2168 Verruconis gallopava, 2155, 2158, 2161, 2163 Verruga peruana, 873, 876 Versant HCV Genotype (LiPA) 2.0, 1606 Versant HCV RNA, 1603–1604 Versant HCV RNA 3.0, 1411, 1610 Versant HIV-1 RNA 3.0, 1411, 1442–1443 VersaTREK blood culture system, 21–22, 23 VersaTREK culture system, 553, 577 VersaTREK fungi, 1948 VersaTREK Myco PZA kit, 1366–1367 VersaTREK Myco susceptibility kit, 1366– 1367 Vertigo, tetracyclines and, 1187 Vesicles, 1942 herpes simplex virus (HSV), 1688 varicella-zoster virus, 1704–1705, 1709 viruses, specimens and methods for detection of, 1406 Vesicomyiasis, 2518 Vesicular fluid, varicella-zoster virus, 1706– 1709, 1712 Vesicular stomatitis virus, 1645 Vesivirus (genus), 1617 Vespidea, 2518 Vestibular toxicity, aminoglycosides and, 1182

n clv

VET-RPLA, 695 Vi antigen, Salmonella, 704 Viannia (subgenus), 2361, 2362 Vibrio, 762–769; see also Vibrionaceae (family); specific species alkaline peptone salt broth for, 327 antimicrobial susceptibilities, 1180, 1186 antimicrobial susceptibility testing, 1318, 1331 clinical significance, 996–997 commercial sources of chromogenic agar media for, 326 identification, 767–768, 994–997 properties of, 763 specimen collection, transport, and storage guidelines, 301–302 sucrose-Teepol-tellurite agar, 345 taxonomy, 762 typing systems, 768–769 Vibrio alginolyticus, 762, 764–765, 767–769, 996–997, 1331 Vibrio carchariae, 766 Vibrio cholerae, 762–769, 996 alkaline peptone salt broth for, 327 alkaline peptone water, 327 antimicrobial susceptibilities, 769, 1186 antimicrobial susceptibility testing, 1331 direct examination, 766–767 El Tor biotype, 763, 767, 1331 epidemiology, transmission, and clinical significance, 762–763 evaluation, interpretation, and reporting of results, 769 identification, 767–768 commercial systems, 768 conventional phenotypic tests, 767–768 molecular methods, 768 toxin detection, 768 in Acanthamoeba, 2389 isolation procedures, 767 O1 serogroup, 126, 762–763, 766–769, 1331 O139 serogroup, 763, 766–769, 1331 phenotypic test results, 764 specimen collection, transport, and storage guidelines, 301–302 taxonomy, 762 TCBS agar for, 345 toxin, 763, 768 typing systems, 768–769 molecular typing, 769 serotyping, 768–769 Vibrio cincinnatiensis, 764, 766, 996–997 Vibrio damsela, see Photobacterium damselae Vibrio extorquens, see Methylobacterium mesophilicum Vibrio fluvialis, 764–765, 768–769, 996 Vibrio furnissii, 764–765, 768, 996 Vibrio harveyi, 764, 766, 769, 997 Vibrio metoecus, 766 Vibrio metschnikovii, 762, 764, 766–769, 996– 997 Vibrio mimicus, 762–765, 767–769, 996 Vibrio navarrensis, 766 Vibrio parahaemolyticus, 762, 764–769, 996 antimicrobial susceptibility testing, 1331 TCBS agar for, 345 Vibrio parahaemolyticus agar, 346 Vibrio parahaemolyticus sucrose agar, 346 Vibrio vulnificus, 764–769, 997, 1331 Vibrionaceae (family), 762–769 antimicrobial susceptibilities, 769 collection, transport, and storage of specimens, 766

clvi

n

SUBJECT INDEX

Vibrionaceae (family) (continued) description of family, 762 direct examination, 766–767 epidemiology, transmission, and clinical significance, 762–766 evaluation, interpretation, and reporting of results, 769 identification, 767–768 commercial systems, 768 conventional phenotypic tests, 767–768 molecular methods, 768 toxin detection, 768 isolation procedures, 767 phenotypic test results, 764 properties of, 763 serolic tests, 769 taxonomy, 762 typing systems, 768–769 molecular typing, 769 serotyping, 768–769 Vidarabine (ara-A) antiviral susceptibility testing, 1916 polyomavirus, 1811 progressive multifocal leukoencephalopathy (PML), 1811 Villose, 1942 VIM type β-lactamases, 1223–1224, 1226, 1383–1384 Vincent’s angina, 300 Viper system, 74 ViraBlot, 1070, 1072 Viral encephalitis, as biothreat agent, 223– 224 Viral gastroenteritis, laboratory tests suggested for, 125 Viral hemorrhagic fever viruses arenaviruses, 1669–1682 filoviruses, 1669–1682 Viral hemorrhagic syndrome, 1647 Viral load assays, 75, 77 Epstein-Barr virus (EBV), 1740, 1742– 1743, 1746–1747 human immunodeficiency virus, 1442– 1443, 1450 Viral replication capacity (RC) assay, human immunodeficiency virus, 1443 Viral transport medium, 1409–1410 ViraTrans medium, 1410 Virdarabine, for herpes simplex virus (HSV), 1689 Virgibacillus, 438 Viridans group streptococci, see Streptococcus, viridans Virion morphology, 1401 ViroChip, 144, 241–242 Virocult medium, 1410 Virohep-A, 1585 Vironostika HTLV-I/II Microelisa system, 1463 ViroSeq HBV genotyping kit, 1920 ViroSeq HIV-1 genotyping system, 1447– 1448, 1450, 1920 ViroSeq HIV-1 integrase genotyping system, 1920 ViroSeqpro-RT system, 1920 Virulence testing, Escherichia coli, 694–695 Virus algorithms for detection and identification of, 1432–1435 cell culture media, 1429–1430 cell cultures, 1424–1429 cell cytotoxicity assays, 1429 cell lines and virus susceptibility profiles, 1428

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 156

centrifugation-enhanced rapid, 1426– 1427 ELVIS, 1429 traditional, 1424, 1426 collection methods and processing of specimens amniotic fluid, 1412 blood, 1412–1413 bone marrow, 1413 cerebrospinal fluid, 1413 dried blood spots, 1413 eye, 1413 genital, 1413–1414 oral, 1414 overview, 1405, 1408–1409 respiratory, 1414–1415 skin, 1415 stool and rectal swabs, 1415 tissue (biopsy or autopsy), 1415–1416 transportation regulations, 1416 urine, 1416 discovery by next-generation sequencing, 242–246 by Sanger sequencing, 242, 243 DNA microarrays for rapid identification, 144–145 enteroviruses, 1536–1546 morphology, 1401 parechoviruses, 1536–1546 reagents, 1422–1423 risk-based classification, 171 specimen collection, transport, and processing, 1405–1417 collection methods and processing of specimens, 1412–1416 specimen collection, 1405, 1408–1409 specimen selection, 1405–1408 specimen storage and processing, 1410– 1412 transport conditions, 1410 transport medium, 1409–1410 stains, 1423–1425 storage methods, 167 subtyping by molecular methods, 76 taxonomy and classification, 1393–1402 character-based descriptors, 1394–1395 criteria, 1394–1397, 1402 future challenges, 1402 human pathogens, 1398–1400, 1402 ICTV classification, 1393–1394 ICTV database, 1402 NCBI database, 1402 sequence-based characters, 1395, 1402 Virus neutralization arboviruses, 1654–1655 poxviruses, 1835–1837 respiratory syncytial virus, 1508 rhinovirus serotyping by, 1557 VirusHunter, 246 VISA, see Vancomycin-intermediate S. aureus Visceral larva migrans Toxocara, 2496 treatment, 2531 Viscerotropic arboviruses, 1647 VisiTect Syphilis, 1069 VISS (vancomycin-intermediate Staphylococcus species), 1294 Vitek 1, 1275 Vitek 2, 32, 428, 453 Acinetobacter, 1279–1280 Advanced Expert System (AES), 1276, 1280 advantages of, 1277

ANC card, 906 anaerobic Gram-negative rods, 977 Clostridium, 954 non-spore-forming, Gram-positive, anaerobic rods, 927 antifungal susceptibility testing in yeasts, 2265 antimicrobial susceptibility testing, 1275– 1280 AST-ST01 card Streptococcus, 1320 Bordetella, 843 coryneform Gram-positive rods, 482 data management system, 1276 Enterobacteriaceae, 1279 Enterococcus, 411, 1278 GN card Burkholderia, 799–800 Pseudomonas, 778 GNI+ Vibrionaceae, 768 Gram-positive card, 364 ID-GNB card, 658–659 Gram-negative nonfermentative rods, 816 Stenotrophomonas maltophilia, 801–802 Vibrionaceae, 768 ID-NH card, 658, 660 Klebsiella pneumoniae, 1279 Neisseria/Haemophilus identification cards, 676, 678 non-spore-forming, anaerobic, Grampositive rods, 930 Pseudomonas aeruginosa, 1279–1280 Smart Carrier Station, 1275–1276 Staphylococcus, 364, 1278–1279, 1289 Streptococcus, 1279 Streptococcus pneumoniae, 1319 YST, 2001 Vitek 2 Compact, 1276 Vitek MS, 48, 72, 583 anaerobic Gram-negative rods, 982 Aspergillus, 2043 Clostridium, 954–955 Haemophilus, 677 IVD, 25, 35, 677 Staphylococcus, 365 Vitek MS Plus, 35 Vitek MS RUO, 35, 677 Vitek MS v2.0, 35–37 Vitox, 672 Vitreous fluid specimen collection, transport, and storage guidelines, 275 fungi, 1945, 1949–1950 Vitros Anti-HBc, 1848 Vitros Anti-HBc IgM, 1848 Vitros Anti-HBs, 1847 Vitros Anti-HCV, 1607–1608 Vitros HBeAg, 1847 Vitros HBsAg, 1847 Vittaforma, 2209–2211, 2330 Vittaforma corneae, 2210, 2212, 2215 ViveST, 1409 Vizion system, Sensititre, 1275 Vogel and Johnson agar, 346 Voges-Proskauer (VP) test, 320 Streptococcus, 391, 395 Volume of sample, 281–282 Volvariella volvacea, 2071, 2075 Vomiting adenoviruses, 1772 arenaviruses, 1673 Balantidium coli, 2417 Blastocystis hominis, 2406 carbapenems, 1177

SUBJECT INDEX chloramphenicol, 1193 clavulanic acid, 1177 Cryptosporidium, 2438 Cyclospora cayetanensis, 2429 Cystoisospora belli, 2428 Ehrlichia chaffeensis, 1138 filoviruses, 1674 herpes B virus, 1697 influenza virus, 1471 macrolides, 1183 nitrofurantoin, 1196 spider envenomation, 2520 sulfonamides, 1192 telavancin, 1189 telithromycin, 1185 tetracyclines, 1187 Trichinella, 2495 Trichuris trichiura, 2459 variola virus, 1830 Voriconazole, 2226–2227 Acanthamoeba, 2395 antifungal susceptibility testing, 2255– 2273 Aspergillus, 2044–2045 Candida, 2004–2005 dermatophytes, 2145 dimorphic fungi, 2121–2122 eumycotic mycetoma fungi, 2181–2182 Fusarium, 2069 hyaline fungi, 2077 melanized fungi, 2167 microsporidia, 2216 mucormycosis, 2097 Naegleria fowleri, 2395 phaeohyphomycosis, 2167 scedosporiosis, 2167 spectrum of activity, 2224, 2226 Talaromyces marneffei, 2048 yeast species, MICs for, 2005 Voriconazole resistance, 2227, 2238–2239 VRE, see Vancomycin-resistant enterococci VRE agar, 346 VRESelect medium, 347 VRSA, see Vancomycin-resistant S. aureus Vulvovaginitis etiologies, usual, 290 Saccharomyces cerevisiae, 1994 VYOO pathogen detection assay, 1381, 1383–1384 VZV, see Varicella-zoster virus VZV tracer, 1707 Wadsworth method, 1342 Wako test, 1978, 1996 WalkAway system, see MicroScan WalkAway system Wallenstein medium, 347 WAP Lab, 50–51 Warts, 1785–1786 Washington, John, 19 WASP (walk-away specimen processor), 49 Wasps, 2518, 2522 Water, superoxidized, 195 Water (tap) agar, 1962 Water beetles, 2521 Water-safety biothreats, 224 Watsonia, 2482 Wautersia, 791 Wautersia eutropha, 791 Wautersiella, 813 Wautersiella falsenii, 624–625, 826–828 Wayson stain, 323 WEE (Western equine encephalitis) virus, 223–224, 1644, 1646–1647, 1651

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 157

Weeksella, 813 Weeksella virosa, 624–625, 826, 828–829 Weight loss Cystoisospora belli, 2428 Enterobius vermicularis, 2454 hookworm, 2456 Leishmania, 2359 Onchocerca volvulus, 2466 Trichuris trichiura, 2459 Weil-Felix febrile agglutination tests, 1129– 1130 Weil-Felix OX-K reaction, 92 Weil’s disease, 1031 Weissella, 422 antimicrobial susceptibilities, 430 clinical significance, 424 identification, 426–427, 429 interpretation of results, 431 Weissella confusa, 424 West Caucasian bat virus, 1633–1634 West Nile virus, 147, 1644–1645, 1647– 1648, 1650, 1652, 1654–1655 Western blot, 101 amebae, 2394 Anaplasma phagocyrophilum, 1143–1144 arenaviruses, 1680 Borrelia, 1046–1047 Ehrlichia chaffeensis, 1142 filoviruses, 1680 herpes B virus, 1697 herpes simplex virus (HSV), 1693 human immunodeficiency virus type 1, 1446–1447 human T-cell lymphotropic viruses (HTLVs), 1461–1465 Lagenidium, 2204 Mycoplasma, 1097–1098 Paragonimus, 2487 parvovirus B19, 1822 poxviruses, 1835, 1837 Pythium insidiosum, 2203 rubella virus, 1530 Toxoplasma gondii, 2380 transmissible spongiform encephalopathies (TSEs), 1863–1864 Treponema pallidum, 1072 Western equine encephalitis (WEE) virus, 223–224, 1644, 1646–1647, 1651 Wet mount sigmoidoscopy material, 2324 sputum specimen, 2331 stool sample concentration, 2317–2318 direct in saline, 2317 urogenital samples, 2326–2327 WGS, see Whole-genome sequencing Wheately trichrome stain, 2314 Wheezing, bocavirus, 1823 Whipple, George Hoyt, 1159 Whipple’s disease, 75, 240, 1159–1165 classical (systemic), 1160 collection, transport, and storage of specimens, 1161–1162 evaluation, interpretation, and reporting of results, 1164–1165 healthy carriers, 1161 isolated organ manifestations, 1160–1161 therapy, 1163–1164 Whipworm, see Trichuris trichiura White Arroyo virus, 1669, 1672 White grain mycetoma, 1967, 2177 White piedra, 1949, 1994, 2148–2149 WHO, see World Health Organization Whole-genome mapping (WGM), 136, 137

n clvii

Whole-genome sequencing (WGS), 141 classification and identification of bacteria, 260 Clostridium, 957 Enterococcus, 413 epidemiological use, 245 workflow for bacterial molecular diagnostics, 244 Whole-genome single nucleotide polymorphism (SNP) typing, 141– 143 Wickerham-Burton method, 2001 Wickerhamomyces, 1988 Wickerhamomyces anomalus, 1985, 1988, 1993, 2000, 2005 Widal test, 92, 704 Widow spiders, 2520 Wilkins-Chalgren anaerobe broth, 347 Williamsia chemotaxonomic and lysosome growth characteristics, 509 clinical significance, 519–520 description of genus, 511 identification, 438 morphologic characteristics, 508 taxonomy, 504–505 Williamsia deligens, 510, 512, 514, 520 Williamsia muralis, 510, 514, 519–520 Winterbottom’s sign, 2366 Wirtz-Conklin spore stain, 323 Wisconsin State Laboratory of Hygiene, 219 Wohlfahrtia, 2330, 2517 Wohlfahrtiimonas chitiniclastica, 632–633, 820, 822 Wolbachia, 1135–1136, 1138–1139, 2535 filarial nematodes, 2465 Mansonella perstans, 2468 Onchocerca volvulus, 2467 Wolbachia persica, 851–852 Wolinella succinogenes, 994, 997, 1013 Wood’s light, 1949, 2136–2138 Working distance, 6 World Health Organization (WHO) Global Outbreak Alert Response Network, 128 Gonococcal Antimicrobial Surveillance Program (GASP), 1322–1323 human immunodeficiency virus disease classification, 1440 LabNet, 1523–1524 outbreak response, 121 polio eradication program, 148 tuberculosis, 1356, 1361 Worm burden estimation, 2323 Wound botulism, 947 Wound debridement, 2518 Wound infection Aeromonas, 754 Alloscardovia omnicolens, 925 anaerobic Gram-negative rods, 967, 969– 970, 972, 974 Arcanobacterium haemolyticum, 479 Bacillus cereus, 443 Bacillus circulans, 443 Bacillus megaterium, 443 Bacillus subtilis, 442 Bacteroides, 971 Bergeyella zoohelcum, 827 Cellulomonas, 479 Citrobacter, 720 Clostridium perfringens, 945 Corynebacterium amycolatum, 479 Corynebacterium jeikeium, 479 Corynebacterium minutissimum, 479

clviii

n

SUBJECT INDEX

Wound infection (continued) Corynebacterium striatum, 479 Corynebacterium tuberculostearicum, 479 Corynebacterium urealyticum, 479 Dermabacter hominis, 479 Dialister, 974 Dysgonomonas, 655 Edwardsiella, 721 Enterococcus, 406, 415 Eubacterium, 924 Filifactor villosus, 924 Finegoldia magna, 911 Fusarium, 2058 Fusobacterium, 973 Gemella, 424 Gordonia, 515 Gram stain and plating medium recommendations, 286 Gram-negative curved bacilli, 997 Gram-positive anaerobic cocci (GPAC), 910–911 Ignavigranum, 425 Kerstersia, 841 Kingella, 655 Leclercia, 722 Microbacterium, 479 Moraxella canis, 814 Mycobacterium, 596–598, 600 Mycoplasma, 1093 Myroides odoratus, 824 Nocardia abscessus, 516 non-spore-forming, anaerobic, Grampositive rods, 923 Paenibacillus alvei, 443 Paenibacillus macerans, 443 Paenibacillus pasadenensis, 443 Paenibacillus vulneris, 443 Parvimonas micra, 911 Pasteurella, 655 Peptoniphilus, 911 Peptostreptococcus anaerobius, 911 Photobacterium damselae, 765 Prevotella, 967, 973 Proteus, 720 Pseudochrobactrum, 824 Pseudomonas, 776 Pseudomonas aeruginosa, 774–775 Robinsoniella peoriensis, 925 Roseomonas, 830 Serratia, 720 Slackia exigua, 925 Solobacterium moorei, 924 Sphingobacterium, 825 Sphingomonas, 826 Staphylococcus, 360 Stenotrophomonas maltophilia, 794 Trueperella pyogenes, 479 Vagococcus, 424 Vibrio alginolyticus, 765, 997 Vibrio harveyi, 766, 997 Vibrio metschnikovii, 766 Vibrio vulnificus, 765 Wound myiasis, 2518 Wound specimen fungi, 1945, 1947–1948 Mycobacterium, 548 specimen collection, transport, and handling, 305 Wright-Giemsa stain, for fungi, 1976 Wright’s Dip Stat stain, 2333, 2335 Wright’s stain for blood parasites, 2335 for parasitology, 2313–2314 WU polyomavirus, 1803–1805, 1810

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : even

Page 158

Wuchereria, arthropod vector of, 2507 Wuchereria bancrofti, 1139, 2461–2465 antigen detection, 2465 clinical significance, 2462, 2464 description of agents, 2461 detection, 2328, 2332 diagnosis, 2464 direct examination, 2464–2465 epidemiology and transmission, 2461–2462 microscopy, 2464–2465 nucleic acid detection, 2465 prevention, 2465 taxonomy, 2461 treatment, 2465, 2531, 2534–2535, 2544 Wukipolyomavirus (genus), 1803 X factor, 667, 672–673, 675 Xanthomonas, 792 Xenodiagnosis, for parasites, 2307, 2365 Xenon vapor (XBO) lamps, 10 Xenopsylla, 2507 Xenopsylla cheopis, 2509–2510 X-linked inhibitor of apoptosis gene (XIAP), 1740 X-linked lymphoproliferative syndrome, 1740, 1747 xMAP system, 59–60, 145, 2042 Xpect Cryptosporidium, 2295, 2441 Xpect Flu A&B, 1474 Xpect Giardia, 2295 Xpect Giardia/Cryptosporidium, 2295, 2441 Xpect Rotavirus, 1624 Xpect RSV, 1504 Xpert EV, 1542 Xpert Flu A, 1477 Xpert HPV assay, 1790 Xpert MRSA, 361, 1293, 1381–1382 Xpert MRSA/SA SSTI test, 361, 1381 Xpert MTB/RIF test, 23, 575–576, 587 Xpert VanA, 1381 Xpert vanA/vanB assay, 407 xTAG gastrointestinal pathogen panel, 690, 1002, 1627, 2320 Cryptosporidium, 2441 Entamoeba histolytica, 2405 xTAG multiplex PCR assay for yeasts, 1997 xTAG respiratory viral panel (RVP), 1478, 1506, 1511, 1543 adenovirus, 1775 parainfluenza virus, 1490 rhinoviruses, 1554–1555 xTAG respiratory viral panel (RVP) fast, 1478, 1506, 1554–1555 xTAG RVP/RVP Fast/RVP Fast v2, 1575 xTAG RVPv1, 1490 Xylose-lactose-Tergitol 4, 347 Xylose-lysine-deoxycholate agar, 347 Xylose-sodium deoxycholate-citrate agar, 347 Yaba monkey tumor virus, 1828–1829, 1831 Yaba-like disease virus, 1828–1829, 1831 Yalow, Rosalyn S., 91 Yaniella, 354 Yarrowia lipolytica, 1985–1986, 2000–2001, 2263 Yatapoxvirus (genus), 1398, 1828–1829, 1831, 1835 Yatapoxviruses, 1828–1837 antigen detection, 1833 clinical significance, 1831 collection, transport, and storage of specimens, 1832 description of agents, 1828 diagnostic tests, 1832

direct detection, 1832–1835 epidemiology and transmission, 1828–1829 evaluation, interpretation, and reporting of results, 1837 identification, 1835 isolation, 1835 microscopy, 1832–1833 nucleic acid detection, 1835 serologic tests, 1837 taxonomy, 1828–1829 Yaws, 1055–1056, 1058, 1061 Yeager’s LIT (liver infusion tryptose) medium, 2315 Yeast(s), 1935, 1984–2006; see also specific species antifungal susceptibilities, 2004–2006 antifungal susceptibility testing broth macrodilution method, 2262 broth microdilution method, 2258–2264 clinical breakpoints, 2263–2264 colorimetric methods, 2264–2265 direct testing on blood samples, 2267 disk diffusion method, 2266 Etest, 2267 flow cytometry methods, 2265–2266 gradient strip testing, 2267 MALDI-TOF MS, 2267 molecular methods, 2267–2268 Neo-Sensitabs, 2267 proteomic methods, 2267 Vitek 2 method, 2265 antigen detection, 1996–1997 ascomycetous, 1938 basidiomycetous, 1937 black, 1939, 2004 clinical significance, 1992–1994 collection, transport, and storage of specimens, 1994 cultural and biochemical characteristics, 1989 description of agents, 1985–1991 ascomycetous yeasts, 1986–1988 basidiomycetous yeasts, 1988–1991 direct examination, 1994–1998 India ink, 1996 KOH, 1995 microscopy, 1995–1996 yeast in tissue sections, 1996 epidemiology and transmission, 1991–1992 evaluation, interpretation, and reporting of results, 2006 identification, 1998–2004 ascospore formation, 2000 carbohydrate assimilation tests, 2001 carbohydrate fermentation tests, 2001 chromogenic agars, 1999 germ tube test, 1999–2000 macroscopic characteristics, 1998 microscopic characteristics, 1998–1999 molecular methods, 2002–2003 morphology studies, 1999 nitrate tests, 2001 phenol oxidase test, 2000 phenotypic systems, 2001 purity of cultures, 1999 rapid identification, 2001–2002 rapid trehalose test, 2001 scheme, 1995 troubleshooting, 2003–2004 urease test, 2000–2001 identification of, 1939 in tissue sections, 1996

SUBJECT INDEX isolation, 1998 MALDI-TOF (MS) identification of, 38 microscopy, 1966 nucleic acid detection, 1997–1998 organisms resembling, 2004 serologic tests, 2004 storage methods, 166 taxonomy, 1984–1985 ascomycetous yeasts, 1984–1985 basidiomycetous yeasts, 1985 typing, 2004 Yeast carbon agar, 1942, 1962 Yeast extract phosphate, 1952 Yeast extract-phosphate agar with ammonia (Smith’s medium), 1962 Yeast Traffic Light PNA FISH kit, 1997 Yellow fever virus, 1644–1645, 1648, 1650, 1654–1655 arthropod vectors, 2507 laboratory tests suggested for, 125 Yellow jackets, 2518 Yellow sac spider, 2521 Yersinia, 738–748 antigen detection, 743–744 antimicrobial susceptibilities, 747–748 biochemical reactivities and characteristics, 740 clinical significance, 741–743 collection, transport, and storage of specimens, 743 description of agents, 739 direct examination, 743–744 epidemiology and transmission, 739, 741 evaluation, interpretation, and reporting of results, 748 identification, 745–746 isolation procedures, 744–745 microscopy, 743 nucleic acid detection, 744 serologic tests, 747 taxonomy, 738–739 typing systems, 746–747 Yersinia aldovae, 738, 739, 740, 748 Yersinia aleksiciae, 738, 740 Yersinia bercovieri, 738, 739, 740 Yersinia enterocolitica antimicrobial susceptibilities, 747–748, 1180, 1183, 1192, 1195 biochemical reactivities and characteristics, 740 cefsulodin-irgasan-novobiocin agar, 332 clinical significance, 742 commercial sources of chromogenic agar media for, 327 description of agents, 739 direct examination, 744 epidemiology and transmission, 741 evaluation, interpretation, and reporting of results, 748 identification, 745–746 isolation procedures, 744–745 nucleic acid detection, 744 serologic tests, 747

MCM 11 Edition X : MCM11$CHSX 03-24-15 06:16:00 PDFd : MCM11X : odd

Page 159

specimen collection, transport, and storage guidelines, 301–302, 743 taxonomy, 738–739 typing systems, 746–747 Yersinia entomophaga, 738, 740 Yersinia frederiksenii, 738, 739, 740, 748 Yersinia intermedia, 738, 739, 740, 748 Yersinia kristensenii, 738, 739, 740 Yersinia massiliensis, 738, 740 Yersinia mollaretii, 738, 739, 740 Yersinia nurmii, 738, 740 Yersinia pekkanenii, 738, 740 Yersinia pestis antimicrobial susceptibilities, 747, 1181 antimicrobial susceptibility testing, 1316, 1324–1325 arthropod vector, 2507 biochemical reactivities and characteristics, 740 biothreat agent, 220, 221 characteristics, 220 clinical significance, 221, 741–742 collection, transport, and storage of specimens, 743 description, 739 differentiation of Francisella from, 852 direct examination, 743–744 epidemiology and transmission, 221, 739, 741 evaluation, interpretation, and reporting of results, 748 genome sequence, 241 identification, 746 isolation procedures, 745 nucleic acid detection, 744 serologic tests, 747 taxonomy, 738–739 typing systems, 747 Wayson stain, 323 Yersinia pseudotuberculosis antimicrobial susceptibilities, 747–748 biochemical reactivities and characteristics, 740 clinical significance, 742 collection, transport, and storage of specimens, 743 description of agents, 739 direct examination, 744 epidemiology and transmission, 741 evaluation, interpretation, and reporting of results, 748 identification, 745–746 isolation procedures, 744–745 nucleic acid detection, 744 serologic tests, 747 taxonomy, 738–739 typing systems, 746–747 Yersinia rohdei, 738, 739, 740, 748 Yersinia ruckeri, 687, 693, 738, 739, 740, 748 Yersinia selective agar base, 347 Yersinia similis, 738, 740 Yimella, 354, 361 YI-S medium, 2315

n clix

Yokenella, 722, 726 Yokenella regensburgei, 718 Zaire ebolavirus, 1670 Zalcitabine, for adenoviruses, 1777 Zanamivir, 1471, 1481, 1886–1887, 1914– 1916 Zanamivir resistance, 1903–1905, 1915, 1917, 1921 Zearalenone, 2189–2191 Zephiran-Trisodium method, of digestion and decontamination, 559 Zeus ELISA HSV-1 and -2 test system, 1693 Zhihengliuella, 354, 361 Zidovudine (AZT) human herpesvirus 8 (HHV-8), 1763 human immunodeficiency virus (HIV), 1870, 1873 human T-cell lymphotropic viruses (HTLVs), 1461 multicentric Castleman’s disease, 1763 Zidovudine resistance, 1896–1898 Ziehl-Neelsen (Z-N) procedure, 321, 2313 Ziehl-Neelsen stain, modified, 2313 Zika virus, 1645 Zimmermannella, 495–496; see also Pseudoclavibacter Zinc tablets, for rhinoviruses, 1558 Zn-PVA, 2311–2312 Zone edge test, 1289 Zone-of-inhibition diameter, 1259–1261, 1265, 1267 Zoonosis Bartonella, 876–877 Brucella, 863 coronaviruses, 1565, 1567, 1569 Cryptosporidium, 2437 Erysipelothrix, 468 Francisella tularensis, 854 hepatitis E virus, 1586, 1588–1589, 1591 herpes B virus, 1687, 1697 leishmaniasis, 2358 Leptospira, 1028–1029 microsporidia, 2212 Plasmodium knowlesi, 2338 poxviruses, 1828–1831 rabies virus, 1633–1634 rotaviruses, 1620 tick-borne infections, 2515 Toxocara, 2496 trematodes, 2479–2490 Zoopagales (order), 2087 Zoospores, 1942, 2199–2200, 2204 Zoster, see Varicella-zoster virus Zoster sine herpete, 1705–1706, 1709 Zygocotylidae, 2290 Zygomycetes, antifungal resistance in, 2243– 2244 Zygomycota (phylum), 1936–1937, 2087 Zygospore, 1937, 1942 Zymodeme analysis, 2402–2404 ZytoFast HPV probes, 1789