Textbook of Practical Microbiology

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Textbook of Practical Microbiology

Subhash Chandra Parija

AHUJA

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Textbook of Practical Microbiology

THE AUTHOR

Dr Subhash Chandra Parija MBBS, MD, PhD, FAMS, FICPath, FABMS, FICAI, FISCD and FIMSA is Director-Professor & Head, Department of Microbiology, in the Jawaharlal Institute of Postgraduate Medical Education & Research, Pondicherry. Prof Parija completed his MBBS at SCB Medical College, Cuttack, Utkal Unioversity, Orissa in 1977. He obtained his MD (1978-81) in Medical Microbiology from the Institute of Medical Sciences, Banaras Hindu University, Varanasi, Uttar Pradesh. Prof Parija did his PhD from University of Madras in the year 1987 for his work on simple diagnostic methods in amoebiasis. Prof Parija, after completion of the MD, began his carrier at the JIPMER, Pondicherry as senior resident in the year 1981. Subsequently He became Professor of Microbiology in the year 1991, and Director-Professor of Microbiology in the year, 2004. During his tenure at JIPMER, Pondicherry, Prof Parija was sent on deputation by Govt of India to set up and establish the departments of Microbiology & initiate an integrated Clinical Laboratory Services at the B P Koirala Institute of Health Sciences, Dharan, Nepal, of which he was the founder head between 1995-98. In recognition of his excellent contribution to the growth and development of the department of Microbiology, as well for the Institute, the B P Koirala Institute of Health Sciences, Dharan, Nepal, conferred the most prestigious BP Koirala Internal Oration Award. Prof Parija was awarded WHO fellowships for study of DNA probes, PCR and other molecular biological methods in the study of parasitic diseases at the University of Aberdeen, UK. Prof. Parija is member of the ICMR task force on intestinal protozoal infections. He is also the member of the Research Advisory Board of BP Koirala Institute of Health Sciences, Dharan, Nepal; Board of MD Examination in Parasitology, Colombo University, Sri Lanka, and the visiting Professor of the College of Medicine & Health Sciences, Sultan Quaboos University, Muscat, Oman. Author of three books “Text Book of Medical Parasitology”, “Stool Microscopy ” and Sputum Microscopy: a Practical Manual”; editor of a book “Review of Parasitic Zoonoses” and two monographs “Immunizing agents for tropics: success, failure and some practical issues” and “Kala-azar: epidemiology, diagnosis and control in Nepal”; Prof Parija also has contributed several chapters for the books, compendium of lectures and proceedings of scientific meetings. Prof Parija has published more than 137 papers both in the National and International journals of repute. Some of his papers are quoted in text books and serial publications. The development of simple, economical and rapid diagnostic tests in serodiagnosis of parasitic diseases such as amoebiasis and cystic echinococcosis is the main field of his research. Prof Parija was the first to demonstrate excretion of hydatid antigen in urine and its detection for diagnosis of cystic echinococcosis. He Developed for the first time the carbon-immunoassay (CIA) and staphylococci adherence test (SAT) as two simple rapid diagnostic methods using a light microscope, and Co-agglutination (Co-A) and CIEP for the detection of antigen in the serum in the patients with amoebic liver abscess and cystic echinococcosis, and for demonstration of antigen in the hydatid fluid and also in the urine for the diagnosis of cystic echinococcosis. All these tests can be used in the field or in less equipped laboratories in the developing countries like India. Prof Parija was first to report the use of LPCB and KOH in the wet mount preparation of stool for detection of intestinal parasites by light microscopy. Prof Parija is the recipient of the most prestigious Dr BC Roy National Award 2003 of the Medical Council of India in recognition of his immense contribution to the development of Medical Microbiology. He was awarded the coveted Dr B P Pandey Memorial Oration Award of the Indian Society for Parasitology, and BK Aikat Oration Award of the Indian Council of Medical Research for his research in diagnosis and epidemiology of parasitic diseases. Other awards include Dr S.R.Memorial Award 2003 of the Bombay Veterinary College Alumni Association, Dr SC Agarwal Oration Award 2001 of the Indian Association of Medical Microbiologists, Major General Saheb Singh Sokhey Award 1992 of the Indian Council of Medical Research, Third Dr Datta Memorial Award 1999 of the Indian Association for the Advancement of Veterinary Parasitology, IAPM (Orissa chapter) Oration Award 1993 of the Indian Association of Pathologists & Microbiologists (Orissa Chapter), Smt Kuntidevi Malhotra Award 1990 of the Indian Association of Pathologists & Microbiologists, Dr S S Misra Memorial Award 1987 of the National Academy of Medical Sciences, Young Scientist Award 1986 of the Indian Association of Medical Microbiologist and Best Scientific Paper Award 1987 of the JIPMER Scientific Society. Professor Parija is the editor-in chief of the online journal Internet Journal of Parasitic Diseases, and member of the editorial editorial boards of various journals both International (Parasitology International, BMC Infectious Diseases, BMC Clinical Pathology and Health Renaissance) and National (Indian Journal of Medical Microbiology, Journal of Veterinary Parasitology, Journal of Parasitic Diseases and Indian Journal of Pathology & Microbiology) . He has been conferred with various fellowships such as FAMS, FICPath, FABMS, FICAI, FISCD and FIMSA by professional bodies. Prof Parija has visited many countries, delivered invited talks in infectious diseases at universities, institutions conferences, seminars etc, and has chaired scientific sessions in the international as well as national conferences. He has guided both MD and PhD students and has been examiner for MBBS, MD and PhD of various universities of India and abroad. Prof Parija was the former Secretary of the Indian Association of Medical Microbiologists and National Vice President of the Indian Association of Pathologists and Microbiologists. He is currently the National Vice President of the Indian Association for Development of Veterinary Parasitologists . He is also life member and member of the executive council of many national and international scientific organizations.

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Textbook of Practical Microbiology

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Textbook of Practical Microbiology

1 / Running Head

Textbook of Practical Microbiology

Dr. Subhash Chandra Parija MD, PhD, FAMS,FICAI, FABMS, FISCD, FIMSA, FICPath

Director-Professor and Head Department of Microbiology Jawaharlal Institute of Postgraduate Medical Education and Research Pondicherry, India.

Ahuja Publishers Bangalore, New Delhi

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Textbook of Practical Microbiology

Textbook of Practical Microbiology Copyright © 2006 Dr. Subhash Chandra Parija

All rights reserved. No part of the publication may be reproduced, stored in retrieval system or transmitted by any means, electronic, mechanical, photocopying or otherwise without the prior writtenpermissionof the publisher.

First Edition : 2006 Printed in India ISBN

Published by Ahuja Publishers

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To my mother

Late Smt. Nishamani Parija

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Textbook of Practical Microbiology

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Contents

Preface

xi

Acknowledgments

xii

UNIT I Microscope and Basic Microbiological Techniques

1

Introduction 1 Compound Microscope 2 Darkground Microscopy 3 Measurement of Microorganisms 4 Hanging drop Preparation 5 Isolation of Pure Cultures

2 3 7 9 11 14

UNIT II Bacterial Staining

19

6 Simple Staining 7 Gram’s Staining 8 Acid Fast Staining 9 Albert’s Staining 10 Capsule Staining 11 Spore Staining 12 Negative Staining

20 23 27 31 34 37 40

UNIT III Cultivation of Bacteria

43

13 14 15 16 17 18 19

44 47 49 51 53 56 59

Media for Routine Cultivation of Bacteria Temperature Requirement for Growth of Bacteria pH Requirement for Growth of Bacteria Oxygen Requirement for Growth of Bacteria Culture of Anaerobic Bacteria Sterilization of Commonly Used Culture Media Antiseptics and Disinfectants

UNIT IV Enzymatic and Biochemical Activities of Bacteria

61

20 21 22 23

62 65 68 71

Catalase Test Oxidase Test Coagulase Test Urease Test

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Textbook of Practical Microbiology

24 25 26 27 28 29 30

Indole Test Methyl Red Test Voges-Proskauer Test Citrate Utilization Test Triple Sugar Iron Agar (TSI) Test Hydrogen Sulphide Test Nitrate Reduction Test

74 76 78 80 82 85 88

UNIT V Antimicrobial Sensitivity Tests

91

31 32 33 34 35

92 95 97 100 102

Kirby-Bauer Method Stoke’s Method Agar Dilution Method Broth Dilution Method Epsilometer Test (E-test)

UNIT VI Immunology

105

Introduction 36 Bacterial Agglutination Test 37 Blood Grouping 38 Latex Agglutination Test 39 Co-agglutination Test 40 Widal Test 41 Weil Felix Test 42 Anti-Streptolysin O (ASLO) Test 43 VDRL Test 44 Radial Immunodiffusion Test 45 Immunoelectrophoresis Test 46 Counter-current Immunoelectrophoresis Test 47 Indirect Haemagglutination Test 48 Immunofluorescence Test 49 Enzyme-linked Immunosorbent Assay

106 108 110 112 114 116 119 121 123 126 128 130 132 135 138

UNIT VII Microbial Genetics and Molecular Techniques

143

Introduction 50 Isolation of Plasmids 51 Polyacrylamide Gel Electrophoresis 52 Isolation of Antibiotic Resistant Mutant 53 Bacterial Conjugation

144 145 148 152 155

Unit VIII Bacteriology

159

54 55 56 57 58 59 60 61 62 63

160 162 164 166 169 172 175 178 181 184

Normal Microbial Flora of the Mouth Normal Microbial Flora of the Throat Normal Microbial Flora of the Skin Identification of Staphylococcus aureus Identification of Streptococcus pneumoniae Identification of b-haemolytic streptococci Identification of Corynebacterium diphtheriae Identification of Lactose Fermenting Enterobacteriacae Identification of Vibrio cholerae Identification of Pseudomonas aeruginosa

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Unit IX Parasitology

187

Introduction 64 Saline Wet Mount of Stool 65 Iodine Wet Mount of Stool 66 LPCB Wet Mount of Stool 67 Acid-fast Staining of Stool Smears 68 Leishman’s staining of Peripheral Blood Smears 69 Concentration of Stool for Parasites 70 Culture of Stool for Entamoeba histolytica

188 189 192 195 198 201 205 208

Unit X Mycology

211

Introduction 71 Cultivation of Fungi 72 Gram’s Staining for Fungi 73 Lactophenol Cotton Blue (LPCB) Wet Mount of Fungi 74 Potassium Hydroxide Wet Mount of Fungi 75 Indian Ink Wet Mount Preparation 76 Slide Culture 77 Germ Tube Test 78 Urease Test 79 Carbohydrate Assimilation Test 80 Carbohydrate Fermentation Test 81 Identification of Common Fungi

212 213 215 217 219 221 223 225 227 229 231 233

Unit XI Virology

239

82 Cultivation of Viruses in the Cell lines 83 Cultivation of Viruses in Embryonated Egg

240 243

Unit XII Microbiology of Water, Milk and Air

247

84 Microbiology of Water 85 Microbiology of Milk 86 Microbiology of Air

248 252 254

Unit XIII Animal Experiments

257

87 Intravenous Inoculation into Mice Tail Vein 88 Collection of Blood from the Marginal Ear Vein of Rabbit 89 Animals and their uses in the Laboratory

258 261 263

Unit XIV Medical Entomology

267

90 Identification of Common Insects

268

Unit XV Common Viva Spots

273

91 Identification of Common Viva Spots

274

Index

295

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Textbook of Practical Microbiology

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Preface

Textbook of Practical Microbiology, is a performance–based text designed for use by students of medicine, microbiology, medical laboratory technology, allied sciences; by laboratory workers and by others who are interested in study of practical microbiology. The intent of the book is to provide recent information and explain in detail the routine diagnostic methods performed in a Microbiology laboratory. Every effort has been made to incorporate all aspects of practical microbiology. A sincere effort is made to provide the essential underlying principles of practical microbiology, to help students to perform various practicals, and to learn and apply the knowledge of practical microbiology in clinical medicine. Textbook of Practical Microbiology consists of 15 learning units. Each unit contains many practical exercises. Each exercise contains learning objectives, theoretical aspects of the practical, principle of the test, and experimental procedure in detail. Important points of the practical experiment are highlighted, possible questions with answers are provided and finally, useful additional informations are provided as box items. The book is profusely illustrated with diagrams, and photomicrographs both black and white, and colour. I owe a special debt of profound gratitude to my mother late Smt. Nishamani Parija and father Shri Mana Govinda Parija without whose encouragement the book would not have been possible. I welcome readers views and suggestions for further improvement of the book in future editions.

Subhash Chandra Parija email: [email protected]

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Textbook of Practical Microbiology

Acknowledgements

I gratefully acknowledge all my colleagues and friends for their valuable advice and assistance in preparation of the manuscript. It is my pleasure to thank my niece Er(Ms) Kukumina Parija, son-in-law Er. Subhasis Ray, nephew Er. Rajkumar Parija, daughter-in-law Mrs Smriti Parija, and daughters Dr. Madhuri Parija and Miss Mayuri Parija for their untiring secretarial help towards the preparation of the manuscript. I am very much thankful to Ahuja Publishers, New Delhi who have been very supportive of this venture.

Subhash Chandra Parija

Textbook of Practical Microbiology

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UNIT

I Microscope and Basic Microbiological Techniques

Introduction Lesson 1 Compound Microscope Lesson 2 Darkground Microscopy Lesson 3 Measurement of Microorganisms Lesson 4 Hanging Drop Preparation Lesson 5 Isolation of Pure Cultures

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UNIT

Introduction Microscope is the instrument, the most important characteristic of microbiology laboratories. The magnification, it provides, enables us to see microorganisms and their structures otherwise invisible to the naked eye. The magnitudes attainable by microscopes range 100X- 400000X. A microscope may be defined as an optical instrument, consisting of a lens or a combination of lenses, for making enlarged or magnified images of minute objects. (Micro: small; scope: to view). Antony Von Leeuwenhoek is considered to be the first person who has seen a micro organism through a simple microscope made by him with a magnification of 270-480 times. He described the size, shape, movements of bacteria, protozoa and algae. These findings were later confirmed after the development of compound microscope by Robert Hooke. The characteristic morphological studies enabled by the discovery of powerful microscopes, helped the scientists to classify microorganisms. Later improvements in the compound microscopes were made and Amici discovered oil immersion lens, which enabled the scientists to study the characteristics more minutely. Microscopes are continuously improved to enable us to have higher magnifications and better resolutions. Microscopes are of two categories: Light or optical microscopes and Electron microscopes depending upon the principle on which the magnification is based. Light microscopy, in which the magnification is obtained by a system of optical lenses uses light waves .The light microscopy includes bright field microscopy, dark-field microscopy, fluorescence microscopy and phase contrast microscopy. On the other hand the electron microscopy uses a beam of electrons in place of light waves for visualization of objects .This includes transmission electron microscopy and scanning electron microscopy.

Textbook of Practical Microbiology

LESSON

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Compound Microscope

LEARNING OBJECTIVES

Microscope stand

After completing this practical you will be able to: 1 Become familiar with the principle, various parts of the compound microscope and its usage in microbiology laboratory. 2 Visualize the cellular morphology from stained slide preparation by using compound microscope.

It is the main framework of the microscope. It consists of : a Main tube. b Body tube. c An arm, which supports the main tube, body tube and the stage. d A substage, and e A foot or base upon which the whole instrument rests.

INTRODUCTION The commonly used microscope in microbiology laboratory is called compound microscope. In compound microscopy, the microscopic field or area observed is brightly lighted and the objects being studied appear dark because they absorb some of the light. Ordinarily microorganisms do not absorb much light but staining them with a dye greatly increases their light absorbing ability resulting in greater contrast and color differentiation. Generally microscopes of this type produce a useful magnification of about 1000x to 2000x. At magnifications greater than 2000x, the image becomes hazy. These microscopes are provided by a coiled filament tungsten lamp. The glowing filaments are prevented from causing glare by focusing their light on the sub-stage condenser, rather than on the object. This is known as Köhler illumination. Closing the aperture of the condenser slightly may aid in detection of certain organisms such as protozoa, fungi, etc. because the light will be hitting the edges of the object at a sharper angle, increasing contrast.

Parts of the compound microscope A microscope mainly consists of 1 Microscope stand. 2 Stage, and 3 Microscope optics.

Main tube: The main tube primarily holds the objective and eyepiece. The eyepiece, also known as ocular piece, is present at the top of the main tube. The eyepiece is fitted loosely into the upper end of the tube. This has a standard diameter so that all the eyepieces are interchangeable. The lower end of the tube contains objectives, which are screwed into what is known as a revolving nosepiece. A number of objectives of lenses of different magnifications are screwed into the nosepiece of the microscope. These objectives can be revolved to increase or decrease the magnification of specimen being examined. Body and arm: The tube is attached to the microscope by the component of the microscope called the body. The body of the microscope and the tube attached to it are supported at the correct height by firm arm, which may also provide a lifting handle for the microscope. Substage: The substage lies immediately below the stage. This holds a condenser lens with an inbuilt diaphragm and a holder for light filter and stops. Foot: The microscope rests firmly on the laboratory bench with the base called foot. This may be U-shaped or rectangular. Stage: A fixed platform with an opening in the center allows for the passage of light from an illuminating source below to the lens system. It provides surface for the placement of a slide over the central opening. Stage can be of fixed or mechanical. Mechanical stage can be moved vertically or horizontally by

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Compound Microscope

means of rack and pinion movement. Stage also contains clips on its surface to hold the slide.

Microscope optics These include objectives, eye pieces, condenser and illuminating source.

Mechanical adjustment of a microscope It is being carried out to focus the specimen examined by the microscope. This adjustment includes coarse and fine focusing adjustments and condenser adjustments.

Coarse and fine focusing adjustments The coarse adjustment is required when focusing the specimens with low power (10x) objectives. It is carried out by rapid and relatively large movements of the stage, which contains the specimen. The fine adjustment is carried out when finer focusing is required by using high power (40x) objectives or oil immersion objectives.

Condenser adjustment Condensers are classified depending on their uses such bright field, dark field, phases contrast, etc. There are four principal types of condensers with respect to correction of optical aberrations, as listed in the table 1-1. The Abbe condenser has two optical lens elements that produce an image of the illuminated field diaphragm that is not sharp and is surrounded by blue and red color at the edges. The next level of condenser correction is split between the aplanatic and achromatic condensers that are corrected exclusively for either spherical (aplanatic) or chromatic (achromatic) optical aberrations. The condenser adjustment system consists of: a Focusing: It is done by moving the condenser up to down. b Adjustment of aperture: It is done by opening or closing iris diaphragm.

Abbe Aplanatic Achromatic Aplanatic achromatic

A good source of light is needed to examine specimens correctly. This may be daylight or electric light. Whatever the source of light, it should fill field of view. It should also fill the whole of the back lens of the objective regularly with light, if not the image will not be clear. While using electric light, a blue filter is placed between the source of illumination and the substage condenser. In some other microscopes, one may find that mirrors are also provided with artificial light. In such case, the flat side of the mirror should be used. When daylight is the natural source of light, the concave mirror should be used without the substage condenser.

PRINCIPLE Magnification The purpose of the microscope is to produce an enlarged, welldefined image of objects too small to be observed with the naked eye. The degree of enlargement is the magnification or magnifying power and it is expressed as the number of times the length, breadth or diameter but not the area of the object is multiplied. The limit of useful magnification is set by the resolving power. Magnification is effected in two stages: the first by the objective lens and the second by the eye-piece lens. The three objectives most commonly used in microbiology laboratory are: (i) A low power objective with focal length 16 mm and magnification 10X. (ii) A high power objective with focal length 4 mm and magnification 40X. (iii) An oil immersion objective with focal length 2 mm and magnification 100X. The total magnification of the microscope can be calculated by multiplying the magnifying power of the objective by that of the eye-piece. With the most powerful lenses, including the 2mm oil immersion objective, the limit of resolution is about 0.2 µm and the greatest useful magnification 1000x or a little less.

Principle involved in the magnification of the object

Table 1-1 Types of condensers Condenser type

The light source

Aberrations corrected Spherical Chromatic -----------X ----------X X X

In biconcave lens, if the object is placed between focal length (f) and 2f, the image is enlarged. This image is real and can be projected on to a screen. If the object is placed between the f and the lens then an enlarged virtual image which cannot be projected on to a screen is produced. In the compound microscope, the object is placed between the f and 2f of the objective lens. The objective produces the primary image. The primary image is real, inverted and magnified. The eyepiece consists of two lenses, a field lens and an eye lens; and a diaphragm between the two lenses. The field lens of the eyepiece

Textbook of Practical Microbiology

brings the real image to focus at the plane of the diaphragm. This is within the focal length (f) of the eye lens. Then the eye lens produces the virtual magnified image that is seen by the eye.

Importance of numerical aperture

5

examine for higher magnification. Focus the 100X objective using the fine adjustment. The condenser may be raised completely upward for obtaining better illumination. 12 Put off illumination and carefully clean all objective lenses and eyepieces with lens paper. 13 Replace the microscope in its box.

The numerical aperture of a microscope objective is a measure of its ability to gather light and resolve fine specimen detail at a fixed object distance. All modern microscope objectives have the numerical aperture value inscribed on the lens barrel, which allows determination of the smallest specimen detail resolvable by the objective and an approximate indication of the depth of field (Box 1-1).

Carefully observe the morphology and colour (after staining) of bacteria present in the smear and also for uniformity in staining, size, shape, Gram’s reaction arrangement.

REQUIREMENTS

RESULTS AND INTERPRETATION

I Equipment Compound light microscope.

The organisms appear dark against a brightly lighted background. The stained smear shows the presence of the bacteria. Sizes of different organisms are summarised in the box 1-1.

OBSERVATIONS

II Reagents Immersion oil and lens wiping paper.

BOX 1-1 TERMINOLOGY III Specimen Stained smear on a glass slide.

PROCEDURE 1 Place the microscope on a firm bench so that it does not vibrate. It should be preferably away from direct sunlight. 2 Put on the illumination when using artificial light. Use the flat side of the mirror to reflect the light up through the condenser when using artificial light. 3 Place the slide to be examined on the stage, making sure the under side of the slide is completely dry. 4 Place the low power objective (10x) in position. Begin examination of the slide with 10x objective. Then to focus the objective, rack the objective carefully down and using the coarse focusing knob and looking it from the side until the lens is near the slide but not touching it. Then while looking through the eyepiece, rack the objective slowly upward, still with the coarse adjustment, until the image comes into view and is sharply focused. 5 Adjust the illumination in such a way that the illumination of the image is optimum. 6 Focus sharply on the specimen using the fine adjustment. 7 Then focus the condenser, for better visualization of the specimen. 8 Examine the specimen, moving it by the mechanical stage. 9 Place the 40x objective in position and examine for higher magnification. Focus the 40x objective using the fine adjustment. The condenser may be raised upward for obtaining better illumination. 10 Place a drop of immersion oil on the smear. 11 Place the oil immersion (100x) objective in position and

Numerical aperture The numerical aperture may be defined as the ratio of the diameter of the lens to its focal length. The greatest possible numerical aperture of a dry lens cannot exceed 1.0. Actually the highest practical numerical aperture of dry and oil immersion lenses is 0.95 and 1.5 respectively. Resolution The limit of useful magnification of a microscope is set by its resolving power. Resolving power is the ability to reveal two closely adjacent structural details as separate and distinct, expressed quantitatively as microscope's limit of resolution i.e. the minimum distance between two visible bodies at which they are seen as separate and not in contact with one another. The greatest resolution in light microscopy is obtained with the shortest wavelength of visible light and an object with maximum numerical aperture. Apochromatic objectives represent high degree of optical perfection used only for critical research and photomicrography due to high expense. Illumination Effective illumination is required for efficient magnification and resolving power. Since the intensity of the daylight is an uncontrolled variable, artificial light from a tungsten lamp is the most commonly used light source in microscopy. The light is passed through the condenser located beneath the stage. The condenser contains two lenses that are necessary to produce a maximum numerical aperture. As the magnification of the lens increases, the distance between the objective lens and slide, called working distance, decreases, whereas the numerical aperture of the objective lens increases.

Compound Microscope

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KEY FACTS 1 2 3 4 5 6

One should remember that proper use and care increases the life of a microscope many fold. Microscope should be kept away from dust, moisture and direct sunlight. After finishing work a cover should be put on the microscope. Care must be taken while handling different parts of microscope. Examination of the slide should always begin with the low power objective (10x). Attempt should never be made to repair microscope by oneself.

BOX 1-2 SIZE OF DIFFERENT ORGANISMS Bacteria Sizes are measured in µm Cocci (spherical shaped bacteria): Size vary from 0.5 µm to 1 µm. Eg. Staphylococcus aureus Bacilli (rod shaped bacteria): Size vary from 1 µm to 10 µm in length and 0.3 µm to 1µm. In breadth. Eg. Bacillus anthracis.

Viruses Sizes are measured in nm. Show variable sizes . Smallest virus measures 20 nm in diameter Eg. Parvovirus Largest virus measures 300 nm in diameter .Eg. Pox virus

Parasites For protozoa sizes are measured in µm Most protozoa are around 50 µm in size Balantidium coli is an exception which measures 100 µm or more in size . For helminths sizes are variable. Sizes are measured ranging from mm to meters Cestodes vary in size from 1mm to several meters in length

Eg: Hymenolepis nana, Taenia saginata, Taenia soluim Nematodes vary in size from 5 mm to even 1 meter in length Eg: Trichinella spiralis, Dracunculus medinesis.

Fungus Sizes are measured in µm Yeast like - appearance 1 2 3 4 5 6 7

Histoplasma capsulatum ___________ Blastomyces dermatitides ___________ Paracoccidioides brasiliensis _______ Sporothrix schenckeii _____________ Candida albicans _________________ Cryptococus neoformans ___________ Chromoblastomycosis _____________

2.3 µm x 3.4 µm 8-15 µm 2-30 µm 2-10 µm 3 µm x 5 µm 4-6 µm 4-12 µm

Mould appearance 1 Aspergillus species ______ 2-5 µm wide hyphae 2 Zygomycetes ____________ 4-5 µm wide hyphae

Spherule like appearance 1 Coccidioides immitis ____ 5-60 µm thick walled spherule 2 Rhinosporidium seeberi __ 200-300 µm sporangia

VIVA 1 Classify microscopes. 2 What is Köhler illumination? Ans Compound microscopes are provided by a coiled filament tungsten lamp. The glowing filaments are prevented from causing glare by focusing their light on the sub-stage condenser, rather than on the object. This is known as Köhler illumination. 3 List the parts of a compound microscope. 4 Define magnification, numerical aperture and resolution. FURTHER READINGS 1 2 3 4

Brooks GF, Butel JS and Morse SA. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd ed. (McGrow Hill, USA.) 2004. Duddington CL. The Microscope. (Museum Press, London) 1964. Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press, London.) 1996. Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds.). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA.) 1997.

Textbook of Practical Microbiology

LESSON

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7

Darkground Microscopy

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to:

1 Take a clean lens wiping paper and carefully clean all the lenses. 2 Place a drop of oil on the depression present on top lens of the condenser. 3 Keep the wet mount preparation on the stage. 4 Raise the condenser slowly so that oil touches the bottom of the slide. 5 Place the low power objective in position in the light path and focus. 6 Using the centering screws adjust the light to fall on the center of the microscopic field. The bright spot of the light must be in the center of the field. If a bright spot is not obtained, slightly raise or lower the condenser to get the bright spot. 7 Place the dry high power objective in the light path, focus and examine the slide. 8 Remove the high power objective from the light path. Place a drop of oil carefully over the cover slip. 9 Now place the oil immersion objective in the light path, focus and observe the slide. 10 Observe the entire field for the motile bacteria, and also observe the type of motility of bacteria. 11 Record the observations in the note book.

1 Observe the motility of microorganisms by dark ground microscopy.

INTRODUCTION Visibility of an object by a microscope depends upon contrast between the object and its background. This can be improved by dark ground microscopy than a compound microscope.

PRINCIPLE In dark-ground microscopy, the object under examination is illuminated not directly but very obliquely (Box 2-1). This is done by opening the aperture of the condenser completely and inserting a funnel stop below the condenser, hence all rays from the condenser are made to pass outside the objective. The rays are reflected or diffracted by the object in the specimen. Hence field appears dark and object is illuminated.

REQUIREMENTS I Equipments Compound light microscope with dark ground condenser. II Reagents Cedar wood oil, lens wiping paper. III Specimen Freshly prepared wet mount preparation of bacteria suspension.

OBSERVATIONS Brightly illuminated motile bacteria are observed under dark background.

RESULTS AND INTERPRETATION The wet mount preparation shows motile bacteria.

Darkground Microscopy

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KEY FACTS 1 Most of the dark ground condensers have fixed focus and must be use with thin slides (1 mm thick) and cover slips (0.1 mm thick). 2 Oil must be used below and above the slide. 3 The wet mount preparation must be thin and should not be dense.

BOX 2-1

PRINCIPLE OF DARK GROUND MICROSCOPY

Dark ground microscopy involves the alteration of microscopic technique rather than the use of dyes or stains to achieve the contrast. By the dark field method, the condenser does not allow light to pass directly through the specimen but directs the light to hit the specimen at an oblique angle. Only light that hits objects, such as microorganisms in the specimens will be deflected upward into the objective lens for visualization. All other light that passes through the specimen will miss the objective, thus making the background a dark field.

VIVA 1 Describe the principle of the dark ground microscopy. 2 List the uses of dark ground microscopy for demonstrating motility. Ans Dark ground microscopy is used commonly for very thin slender bacteria such as spirochetes, which are not visible under ordinary illumination. The motility of spirochetes such as treponemes is clearly seen in a dark ground microscope. Dark ground microscope also is commonly used for demonstrating motility of the trophozoites of protozoa such as Trichomonas vaginalis, Entamoeba histolytica, etc.

FURTHER READINGS 1 2 3 4

Brooks GF, Butel JS and Morse SA. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd ed. (McGrow Hill, USA.) 2004. Duddington CL. The Microscope. (Museum Press, London) 1964. Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press, London.) 1996. Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds.). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA.) 1997.

Textbook of Practical Microbiology

LESSON

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9

Measurement of Microorganisms

LEARNING OBJECTIVES After completing this practical you will 1 Measure the size of bacteria using a microscope. 2 Become familiar with the calibration of an ocular micrometer.

counting the number of spaces occupied by the organism and second by multiplying this number by the calculated calibration factor for one ocular division.

REQUIREMENTS

INTRODUCTION

I Equipments Compound light microscope with ocular micrometer and stage micrometer

Measurement of size of bacteria in a microscope can be done by micrometry.

II Reagents Cedar wood oil and lens wiping paper.

PRINCIPLE

III Specimen Heat fixed smear of Escherichia coli

First the diameter of the microscopic field must be established by means of optic devices, namely, an ocular micrometer and a stage micrometer. The ocular micrometer is placed on a circular shelf inside the eyepiece, which contains graduations on its surface. The distance between these graduations will vary depending on the objective being used, which determines the size of the field. This distance is determined by using a stage micrometer, which also contains graduations that are 0.01 mm apart. The calibration procedure for the ocular micrometer requires that the graduations on both micrometers be superimposed on each other. This is accomplished by rotating the ocular lens. A determination is then made of the number of ocular divisions per known distance on the stage micrometer. Then the calibration factor for one ocular division is calculated by the formula: Known distance between two lines on stage micrometer One division on ocular micrometer = —————————————— Number of divisions on ocular micrometer

Then the size of microorganisms can be determined, first by

PROCEDURE 1 Carefully place the ocular micrometer into the eyepiece. 2 Place the stage micrometer on the microscope stage and center it over the illumination source. 3 With the stage micrometer in clear focus under the lowpower objective, slowly rotate the eyepiece to superimpose the ocular micrometer graduations over those of the stage micrometer. 4 Add a drop of immersion oil to the stage micrometer; bring the oil immersion objective into position and focus. 5 Move the mechanical stage so that a line of the stage micrometer coincides with the line of the ocular micrometer at one end. Fix another line on the ocular micrometer that coincides with a line on the stage micrometer. 6 Determine the distance on the stage micrometer (number of divisions X 0.01 mm) and the corresponding number of divisions on ocular micrometer. 7 Determine the value of the calibration factor for the oil immersion objective. 8 Remove the stage micrometer.

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Measurement of Microorganisms

9 Calculate the number of ocular divisions occupied by each of three separate bacteria and determine the average. 10 Determine the size determined by multiplying the average by calibration factor. 11 Record the observations in the note book.

QUALITY CONTROL A stained smear of standard bacterial strain (e.g., Klebsiella pneumoniae) with known size is examined for comparison.

OBSERVATIONS 1 Observe the divisions on ocular micrometer. 2 Observe the calibration of ocular micrometer of oil immersion objective. 3 Observe the calibration factor. 4 Determine the size by multiplying the average with calibration factor.

RESULTS AND INTERPRETATION The size of E. coli is 2.5 µm in length and 0.6 µm in breadth.

KEY FACTS 1 Carefully clean the eyepiece and objective lenses before starting experiment. 2 Expert technician help must be taken for inserting ocular micrometer. 3 There must be proper coincidence between the lines of stage micrometer and ocular micrometer.

VIVA 1 Explain the method for the measurement of the size of a microorganism by microscopy. 2 Describe the principle involved in the measurement of the size of microorganisms by microscopy.

FURTHER READINGS 1 2 3 4

Brooks GF, Butel JS and Morse SA. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd ed. (McGrow Hill, USA.) 2004. Duddington CL. The Microscope. (Museum Press, London) 1964. Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press, London.) 1996. Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds.). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA.) 1997.

Textbook of Practical Microbiology

LESSON

4

11

Hanging Drop Preparation

LEARNING OBJECTIVES After completing this practical you will be able to:

Spirochaetes are the examples of bacteria which are motile but without presence of any external flagella . The list of motile and non-motile bacteria are summarized in the table 4-1.

1 Observe motility, and also observe natural sizes and shapes of the cells by hanging drop preparation.

REQUIREMENTS INTRODUCTION Hanging drop preparation is one of the easiest method to observe motility in a clinical microbiological laboratory. This is carried out by putting a loop full of bacterial suspension on the cover slip and placing it over a cavity slide and observing it under a microscope. Advantage of this method is that by this method live bacteria can be observed. Examination of living organisms is useful to observe cell activities, viz. motility, binary fission, and also to observe natural sizes and shapes of the cells. Motility of bacteria can also be demonstrated by i. Craigie ’s tube method , ii. swarming of the bacteria on a non inhibitory medium ( e.g, blood agar ) and iii. by dark ground microscopy. Capillary tube method is a useful method for demonstrating the motility of anaerobic bacteria (Box 4-1).

PRINCIPLE Microorganisms such as bacteria, because of their small size and a refractive index that closely approximates that of water, do not lend themselves readily to microscopic examination in a living, unstained state. Bacteria are motile due to the presence of flagella .Depending on the location of the attachment of the flagella , bacteria can be classified as i. monotrichous (single polar flagellum at one end) e.g, Vibrio cholerae , ii. amphitrichous (single flagellum at both the ends) e.g Pseudomonas aeruginosa, iii. lophotrichous (tufts of flagella at one or both ends ) e.g, Spirilla, and iv.peritrichous ( fagella arranged all around the cell) e.g, Escherichia coli, Salmonella sp , etc .

I Equipments Compound light microscope. II Reagents and glass wares Normal saline, inoculating loop wire, staining tray, cavity slides and cover slips. III Specimen Log phase broth culture of E. coli

PROCEDURE 1 Take a clean grease free cavity slide. 2 Take a clean coverslip, apply paraffin to four corners of coverslip. 3 Place a drop of broth culture on the coverslip with the help of inoculating loop. 4 Place the cavity slide (cavity down) over the coverslip so that the drop is placed in center. 5 Invert the slide, and observe under microscope. 6 First observe under low power (10x), locate the edge of the drop, shift the focus to high power (40x) and observe. 7 Record the observation in a notebook.

QUALITY CONTROL The test wet mount preparation is compared with known motile culture of P. aeruginosa for appropriate details such as motility, and natural sizes and shapes of the bacteria.

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Hanging drop Preparation

OBSERVATIONS The bacteria showing motility are demonstrated in the hanging drop preparation. Note: It is important to differentiate active motility from brownian movement. Brownian movement is not true motility, instead it is exibited due to movement of organism as a result of

their collision with water molecules. This movement is usually seen around the axis of bacteria.

RESULTS AND INTERPRETATION The wet mount preparation shows motile bacteria.

BOX 4-1 DEMONSTRATING MOTILITY OF ANAEROBIC BACTERIA Capillary tube method is a useful method for demonstrating the motility of anaerobic organisms. A Robertson’s cooked meat medium or thioglycollate broth culture, inoculated and incubated with anaerobic bacteria is taken. A broth culture of anaerobic bacteria is prepared and is made to fill into the open capillary tube by capillary action. Once the tube is filled up, it is sealed at both ends with clay to maintain anaerobiosis. The tube is then observed, under the high power objective of the light microscope for the presence of motile anaerobic bacteria.

Table 4-1 Motile and non-motile bacteria Motile bacteria

Non motile bacteria

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29

1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20 21 22 23 24 25 26 27 28 29 30 31 32 33 34 35

Other Bacillus species excluding Bacillus anthracis Clostridium tetani Clostridium botulinum Escherichia coli Serratia Proteus Salmonella except S.Gallinarum and S. Pullorum Vibrio cholerae Aeromonas Pleisiomonas Campylobacter Helicobacter Mobiluncus Pseudomonas Burkholderia pseudomallei Burkholderia cepacia Yersinia enterocolitica Ligionella pneumophila Bordetella parapertusis Bordetella bronchiseptica Spirochaetes Treponema Borrelia Leptospira Listeria Spirillum minus Chromobacterium violaceum Eikenella corrodens Alcaligenes faecalis

Staphylococci Micrococci Streptococci Pneumococcus Meningococcus Corynebacterium diphtheriae Bacillus anthracis Clostridium perfringens Lactobacillus Bifidobacterium Propionibacterium Bacteroides Fusobacterium Leptotrichia Klebsiella Shigella Burkholderia mallei Yersinia spp other than Yersinia enterocolitica Pasteurella multocida Francisella tularensis Haemophilus influenzae Moraxella Gardenella Bordetella pertusis Brucella Mycobacterium Mycoplasma Actinomyces Erysipelothrix Streptobacillus Flavobacterium Calymnotobacterium Cardiobacterium Rickettsia Chlamydiae

Textbook of Practical Microbiology

KEY FACTS 1 Always make a thin emulsion of bacterial suspension on the glass slide. 2 Drop should not touch the surface of cavity. 3 Observe the preparation immediately.

VIVA 1 2 3 4

List other methods to confirm motility of the bacteria. Mention the bacterial appendage responsible for motility of the bacteria. List motile and non-motile bacteria. List a non-flagellated motile bacterium.

FURTHER READINGS 1 2 3 4

Brooks GF, Butel JS and Morse SA. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd ed. (McGrow Hill, USA.) 2004. Duddington CL. The Microscope. (Museum Press, London) 1964. Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press, London.) 1996. Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds.). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA.) 1997.

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LESSON

5

Isolation of Pure Cultures

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know various procedures for the separation and isolation of the discrete colonies from a mixed culture or a clinical specimen. 2 Perform the streak plate inoculation procedure. 3 Perform the spread plate inoculation procedure. 4 Perform the pour plate inoculation procedure

INTRODUCTION The microbial population in our environment is large and complex. Many different microbial species normally inhabit various parts of our body such as oral cavity, intestinal tract, skin , etc. These microorganisms may be present in extremely large quantities. Environment, air, soil and water also consist of mixed populations of bacteria and other organisms. Mixed culture is a culture that contains more than one kind of microorganisms, whereas pure culture contains only one kind of microorganisms. Isolation of pure cultures is very important in a clinical microbiology laboratory. Most studies and tests of the physiological, immunological and other characters of bacteria are valid only when made on a pure culture ( Box 5-1) . The preparation of pure culture involves not only the isolation of a given microorganism from a mixed natural population but also the maintenance of the isolated individual and its progeny in an artificial environment to which the access of other microorganisms is prevented. Obtaining well discreet colonies and isolation of a pure culture initially require the number of organisms in the inoculum be reduced. Pure culture isolation methods include streak plate technique, spread plate technique and pour plate technique.

PRINCIPLE Obtaining well discreet colonies and isolation of a pure culture initially require the number of organisms in the inoculum be reduced. The resulting diminution of the population size ensures that, following inoculation, individual cells will be sufficiently far apart on the surface of the agar medium. Streak plate method, a rapid qualitative isolation method, involves spreading a loop full of culture over the surface of an agar plate. Spread plate method depends on spread of the bacterial colonies over the surface of a solid agar medium with a sterile, L- shaped bent gloss rod while the Petri dish is spun on a turntable. Then the plates are incubated at 37°C for 48-72 hours. The well discrete colonies grown on the medium are carefully transferred in to a fresh medium by any of pure culture method. This confirms the purity of isolated colony. Pour plate method is a rapid quantitative isolation method. The method requires a serial dilution of the mixed culture by means of a pipette. This method involves pouring a specimen of molten agar (at 45°C) mixture in a Petri dish; and allowing it to solidify and incubating at 37°C for 48-72 hours. Careful transfer of well discrete colonies in to a fresh medium by any of pure culture method (generally streak plate method) confirms the purity of isolated colony.

REQUIREMENTS I Equipment and labwares Bunsen flame, inoculating loop and marker pen (For all methods). Petri plates, turntable, L-shape glass rod and sterile beaker (Streak plate method). Water bath, thermometer, test tube rack, sterile pipettes, mechanical pipetting device, sterile Petri dishes and disinfectant, (Pour plate method). II Reagents Sterile trypticase soy agar plates (For all methods). 70% ethanol (Spread plate method). Molten trypticase soy agar (Pour plate method).

Textbook of Practical Microbiology

III Specimen Mixed bacterial culture (24-48 hour nutrient broth cultures), exudates, stool and other clinical specimens.

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colony to a fresh agar plate with the help of sterile loop. Repeat the spread plate procedure. 11 After incubation confirm the purity of isolated colony.

For pour plate method PROCEDURE For streak plate method 1 Take a sterile trypticase soy agar plate and label on the back of the agar surface 1, 2, 3, 4 at four corners, approximately 1 cm away from the edge of the plate. 2 Take an inoculating loop and flame to deep red and cool. 3 Charge the inoculating loop with the specimen to be cultured. 4 Transfer the loop full of the specimen (control and test) on to the surface of a well-dried agar plate (area 1), then spread over a small area at the periphery near the flame. 5 Re flame and cool the loop. 6 Turn the Petri dish 90° and touch the loop to a corner of the culture in the area 1 and drag it several times across the agar in area 2. 7 Repeat the same in area 3 and area 4. Note: Flaming of loop can be decided based upon the turbidity of the sample. 8 Incubate all plates in an inverted position for up to 48-72 hours at 37°C. 9 Carefully observe the colony morphology of different colonies. 10 Transfer aseptically the single well-isolated colony on to the surface of other agar plate. 11 Re flame and cool the loop and repeat the same for all colony types. 12 Incubate all plates at 37°C for 48- 72 hours.

1 Liquify 2 agar deep tubes, each containing approximately 20 ml of medium, in an autoclave or by boiling and cool the molten agar tubes and maintain in a water bath at 45°C. 2 Take 2 sterile test tubes and label as test and control. 3 Then add mixed cultures to appropriate tube. 4 Take two sterile dry Petri plates and label as test and control. 5 Remove a tube from the water bath and check the temperature of molten agar medium (45°C). 6 Wipe the outside surface dry with a tissue paper, and aseptically add 1 ml of test culture, mix the contents well by rolling the tube between the two hands carefully and pour the mixture in to test Petri plate. Allow the agar to solidify. 7 Take the second tube, wipe the outer surface aseptically, add 1 ml of control culture, mix the contents well and pour the mixture in to control Petri plate. Allow the agar to solidify. 8 After solidifying the agar, incubate the plates in inverted position at 37°C for 48-72 hours. 9 Select a well-discreet colony from each type and aseptically transfer to a fresh separate agar plate and repeat the streak plate method and incubate the plates at 37°C for 48-72 hours. 10 After incubation confirm the purity of isolated organism.

QUALITY CONTROL Known mixed culture containing standard strains.

For spread plate method

OBSERVATIONS

1 Take 2 trypticase soy agar plates and mark as test and control. 2 Take L shaped 2 glass rods, dip the lower bent portion into 70% ethanol. 3 With a sterile pipette, transfer 0.1 ml of culture from tube 1 to the first agar plate that has been placed on the turntable. Replace the cover. 4 Remove the glass rod from ethanol and pass it through the Bunsen burner flame, with the bent portion of the rod pointing downward to prevent the burning alcohol from running down to arm. Allow the alcohol to burn off the rod completely. Cool the rod for 10-15 seconds. 5 Remove the Petri dish cover and spin the turntable. 6 While the turntable is spinning, lightly touch the sterile bent rod to the surface of the agar and move it back and forth. 7 Remove the glass rod, dip in alcohol and re flame. 8 Stop the turntable. Replace the cover on agar plate. 9 Remove the agar plate from turntable, and incubate in an inverted position at 37°C for 48-72 hours. 10 Carefully observe the colonies and transfer a well-discreet

For streak plate method 1 Observe all the plates for discrete colonies after 24, 48 and 72 hours after first incubation. 2 Search the entire plate for colonies present outside the streak lines, if so discard the plate and repeat the experiment.

For spread plate method 1 Observe the control and test plates after 24, 48 and 72 hours after first incubation. 2 Note the results and colony morphology of different colonies.

For pour plate method 1 Check the control plates for isolated similar looking colonies only.

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Isolation of Pure Cultures

2 Note the colony characters of well- discrete colony of all types of colonies (after both incubations). 3 Preserve the colonies for further characterization.

RESULTS AND INTERPRETATION For streak plate method 1 Check the control plates for isolated similar looking colonies only. 2 Note the colony characters of well- discrete colony of all types of colonies, which are present only on streak lines (after both incubations). 3 Preserve the colonies for further characterization.

For spread plate method 1 Check the control plates for isolated same colonies only. 2 Note the colony characters of well- discrete colony of all types of colonies (after both incubations). 3 Preserve the colonies for further characterization.

For pour plate method 1 Check the control plates for isolated same colonies only. 2 Note the colony characters of well- discrete colony of all types of colonies (after both incubations) from the test plates. 3 Preserve the colonies for further characterization.

VIVA 1 Define pure culture. 2 What are the uses of pure cultures in a microbiology laboratory? Ans. In the microbiology laboratory pure culture are used to: Isolate bacteria in pure culture. Demonstrate their properties. Obtain sufficient growth for preparation of antigens and for other tests. Type the isolates. Determine antibiotic sensitivity pattern. Estimate viable counts. Maintain stock culture. 3 List the selective methods used in the isolation of pure cultures. Ans. Selective methods which help in the isolation of pure culture include: Chemical methods Use of a special carbon or nitrogen source Use of dilute media Use of inhibitory or toxic chemicals Physical methods Heat treatment Incubation temperature pH of the medium Biological methods Animal experiments 4 List the pure culture methods. Write their principles.

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BOX 5-1 TERMINOLOGY A culture that contains only one type of microorganisms is known as pure culture. Pure culture is also called axenic culture. Separation of particular microorganism from a mixed population that exists in nature is called isolation. The descendants of a single isolation in pure culture comprise a strain. If a strain is developed from a single parent cell, it is termed as a clone.

KEY FACTS 1 Surfaces of agar plates must be dry. 2 Whole procedure must be done near the flame and inside a laminar flow inoculation chamber. 3 The loop should be properly sterilized and cooled. 4 Agar plates must be incubated at inverted position. 5 The exposure time of agar surface to the atmosphere should be minimized. 7 Loop should not touch the wall of the Petri dish any time. 8 Loop should not enter any other area except the specified in each step. 9 Contamination while transferring material from one tube to another should be avoided. 10 Occurrence of air bubbles while pouring the medium-culture mixture into plates should be avoided

FURTHER READINGS 1 2 3 4

Brooks GF, Butel JS and Morse SA. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd ed. (McGrow Hill, USA.) 2004. Duddington CL. The Microscope. (Museum Press, London) 1964. Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press, London.) 1996. Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds.). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA.) 1997.

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UNIT

II Bacterial Staining

Lesson 6

Simple Staining

Lesson 7

Gram’s Staining

Lesson 8

Acid Fast Staining

Lesson 9

Albert’s Staining

Lesson 10 Capsule Staining Lesson 11 Spore Staining Lesson 12 Negative Staining

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LESSON

6

Simple Staining

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Stain smears by simple staining method. 2 Compare various morphological forms and arrangement of bacterial cells.

Preparation of polychrome methylene blue: This is prepared by allowing Loeffler’s methylene blue to ‘ripen’ slowly and shaking it at intervals to aerate the contents thoroughly. The slow oxidation of the methylene blue forms a violet compound that gives the stain its polychrome properties. The ripening takes 12 months or more to complete. The stain maybe ripened quickly by addition of 1% potassium carbonate to the stain. This stain gives the acidic cell structures of the bacteria, a red-purple hue.

INTRODUCTION Simple staining employs staining of bacterial smears with a single staining reagent. The commonly used simple stains are the basic stains such as methylene blue, crystal violet and carbol fuchsin. They provide good colour contrast and impart the same colour to the stained bacteria (Box 6-1).

PRINCIPLE In simple staining, the bacterial smear is stained with a single reagent. Basic stains with chromophore are used in simple staining methods.Bacterial nucleic acids and certain cell wall components carry a negative charge that strongly attracts and binds to the cationic positively charged chromogen, and imparts same colour to all bacteria.

REQUIREMENTS I Equipments Compound light microscope. II Reagents and glass wares Loeffler’s methylene blue, polychrome methylene blue, carbol fuchsin, water, microscopic glass slides and Bunsen burner. Preparation of Loeffler’s methylene blue stain: This is prepared by dissolving 30 ml of saturated solution of methylene blue in alcohol, in 100 ml of distilled water containing 0.1 ml of potassium hydroxide (0.01%).

III Specimen Pus, CSF, aspirations.

PROCEDURE 1 Make a thin exudate smear on a slide. 2 Heat fixes the smear by passing the slide 2–3 times gently over the Bunsen flame with the smear side up. 3 Pour Loeffler’s methylene blue over the smear and allow it to stand for 3 minutes. 4 Wash the stained smear with water and air dry it. 5 Observe the smear first under low power (10x) objective, and then under oil immersion (100x) objective. 6 Record the observations in the note book.

QUALITY CONTROL The morphology of the cellular components and bacteria in the known methylene blue stained smear (positive control smears) are compared with that of the blue stained structures in the test smear.

OBSERVATIONS 1 Blue coloured spherical cells, approximately 0.5–1 µm size seen arranged in clusters (Fig. 6-1).

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2 Blue coloured elongated rod shaped cells, approximately 3–5 µm length and 0.2–1.5 µm breadth seen. 3 Blue coloured round cells with darkly stained multilobed nucleus in pus cells are also seen.

RESULTS AND INTERPRETATION The stained smear contains mixture of methylene blue stained cocci in clusters (Fig. 6-1), blue coloured bacilli and the blue coloured multilobed pus cells.

FIGURE 6-1. Methylene blue stained smear of Staphylococcus aureus, x 1000.

BOX 6-1 SIMPLE STAINS AND THEIR USES IN MICROBIOLOGY LABORATORY Loeffler’s methylene blue: It demonstrates morphology of polymorphs, lymphocytes and other cells more clearly. Polychrome methylene blue: It demonstrates the presence of bacteria in smear or wet mount preparation in addition to the morphology of polymorphs and lymphocytes. It is also employed to demonstrate the acidic capsular material of the anthrax bacilli as seen in the Mc Fadyean reaction.

KEY FACTS 1. Methylene blue staining is an example of simple staining. 2. Simple staining method uses a single reagent that imparts the same colour to all bacteria and cellular organelles. 3. Simple staining method shows the presence of bacteria and cellular contents in exudate smears and also in wet mount preparations. 4. Loeffler’s methylene blue shows the characteristic morphology of polymorphs, lymphocytes and other cells more clearly than stronger stains such as Gram’s stain. 5. Smears are required to be stained for 3 minutes and tissues for 5 minutes by simple Loeffler’s methylene blue stain.

VIVA 1 Describe differences between a simple stain and a differential stain. Ans. Simple stains are those that contain only a single-coloured dye, and these when used impart the same colour to all the bacteria in a stained smear. It shows the morphology of leucocytes and other cells more clearly. Methylene blue is an example of a simple stain. Differential stains are those stains that contain two different coloured dyes, hence they impart different colours to different bacteria or bacterial structures, thus differentiating different groups of bacteria. But morphology of leucocytes and other cells are stained less clearly by differential stains. Gram’s stain and acid-fast stain are two examples of differential stains. 2 Name other simple stains. Ans. Loeffler’s methylene blue, polychrome methylene blue and dilute carbol fuchsin are the examples of other simple stains. 3 Mention the uses of methylene blue staining in a diagnostic microbiology laboratory. Ans. Methylene blue staining is used to show a. The presence of bacteria in a smear or wet mount preparation. b. The nature of cellular contents in exudates, and c. The characteristic morphology of polymorphs, lymphocytes and other cells more clearly than the Gram stain. 4 List the uses of polychrome methylene blue staining. Ans. Polychrome methylene blue staining is employed for demonstration of the acidic capsular structure of anthrax bacillus, as shown in the Mc Fadyean reaction.

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Simple Staining

FURTHER READINGS 1 Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press.) 1996. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5 th ed. (Lippincott Williams and Wilkins.) 1997. 4 WHO. Guidelines on Standard Operating Procedures for Microbiology. Chapter 4: Staining Techniques. 5 WHO. Manual of Basic Techniques for a Health Laboratory, 1980.

Textbook of Practical Microbiology

LESSON

7

23

Gram’s Staining

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Stain smears by Gram’s staining method. 2 Differentiate between Gram positive and Gram negative bacteria.

INTRODUCTION Gram’s stain was originally devised by histologist Christian Gram (1884) as a method of staining bacteria in tissues. Gram positive bacteria stain purple, while Gram negative bacteria stain pink when subjected to Gram staining. They stain differently because of the fundamental differences in the structure of their cell walls. Approximately, 60%–90% of the Gram positive bacterial cell wall is made up of peptidoglycan and interwoven teichoic acid, while only 10%–20% of Gram negative bacterial cell wall is composed of peptidoglycan, forming 2–3 layers. The Gram negative cell wall is also surrounded by an outer membrane composed of phospholipids, lipopolysaccharide, lipoproteins and other proteins.

PRINCIPLE The Gram reaction is dependent on permeability of the bacterial cell wall and cytoplasmic membrane, to the dye-iodine complex. In Gram positive bacteria, the crystal violet dye-iodine complex combines to form a larger molecule which precipitates within the cell. Also the alcohol/acetone mixture which act as decolourizing agent, cause dehydration of the multi-layered peptidoglycan of the cell wall.This causes decreasing of the space between the molecules causing the cell wall to trap the crystal violet iodine complex within the cell. Hence the Gram positive bacteria do not get decolourised and retain primary dye appearing violet. Also, Gram positive bacteria have more acidic protoplasm and hence bind to the basic dye more firmly.

In the case of Gram-negative bacteria, the alcohol, being a lipid solvent, dissolves the outer lipopolysaccharide membrane of the cell wall and also damages the cytoplasmic membrane to which the peptidoglycan is attached. As a result, the dye-iodine complex is not retained within the cell and permeates out of it during the process of decolourisation. Hence when a counter stain is added, they take up the colour of the stain and appear pink. Different modifications of Gram’s stainings are summarized in the box7-1.

REQUIREMENTS I Equipments Compound light microscope.

II Reagents and glass wares Bunsen flame and loop wire, clean grease free slides, marker pen, methyl violet (basic dye), Gram’s iodine (mordant), 95% ethanol (decolourising agent), and 1% safranine or dilute carbol fuchsin (counter stain). Preparation of methyl violet stain: This is prepared by dissolving two solutions: solutions A and B. Solution A is made by dissolving completely 2 grams of crystal violet in 120 ml of ethyl alcohol. Solution B is made by dissolving 0.8 grams of ammonium oxalate in the above solution A. Then both solution A and B are mixed well. Preparation of Gram’s iodine: This is made by first dissolving 20 grams of potassium iodide in 250 ml of distilled water and then 10 grams of iodine is further added to it with dissolution. Then 750 ml of distilled water is added and final volume is made up to 1000 ml. Preparation of 1% safranine: This is prepared by dissolving 1 gram of safranine in 100 ml of distilled water.

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Gram’s Staining

III Specimen Preparation of bacterial smear: (From liquid culture) 1 Take clean, and grease free glass slides for making the smears. Note: Wipe the slide with a cotton swab dipped in alcohol and then holding its end with forceps roast it free from grease by passing 6 – 12 times through a blue Bunsen flame. 2 Take one or two loopfuls of the bacterial cell suspension and place on the clean slide with a bacteriological loop. 3 Then with circular movement of the loop, spread the cell suspension into a thin area. 4 Allow the smear to air dry. 5 Heat fix the smear while holding the slide at one end, and by quickly passing the smear over the flame of Bunsen burner two to three times. Preparation of bacterial smear: (From solid media) 1 Take clean and grease free glass slides for making the smears. 2 Take a loopful of 0.85% saline and place it on the centre of the slide. 3 With a straight wire touch the surface of a well isolated colony from the solid media and emulsify in the saline drop forming a thin film. 4 Allow the smear to air dry. 5 Heat fix the smear while holding the slide at one end, and by quickly passing the smear over the flame of Bunsen burner two to three times.

PROCEDURE 1 Cover the smear with the methyl violet (basic dye). Allow it to stand for one minute. 2 Rinse the smear gently under tap water. 3 Cover the smear with Gram’s iodine (mordant) and allow it to stand for one minute. 4 Rinse the smear again gently under tap water. 5 Decolourise the smear with 95% alcohol.

Note: Decolourisation is a critical procedure. Hold the slide with hand on one corner in a slant position and add drop by drop of absolute alcohol till a faint colour comes out of the smear. 6 Rinse the smear again gently under tap water. 7 Cover the smear with dilute carbol fuchsin (counter stain) for 30 seconds to 1 minute. 8 Rinse the smear again gently under tap water and air dry it. 9 Observe the smear first under low power (10x) objective, and then under oil immersion (100x) objective. 10 Record the observations in the note book. Note: List of Gram positive and Gram negative bacteria are summarized in the table 7-1.

QUALITY CONTROL On the same slide, smears of Staphylococcus aureus (Gram positive bacteria) and Escherichia coli (Gram negative bacteria) are made. The slide with control and test smears is stained by Gram’s staining. The appearance of purple coloured Gram positive bacteria and pink coloured Gram negative bacteria in the control smears indicate proper staining technique and stained test smear is compared with it.

OBSERVATION Presence of 0.5–1 µm sized purple coloured spherical bacteria arranged in clusters ( Fig. 7-1) and 1–2 mm sized pink coloured rod shaped bacteria seen ( Fig. 7-2).

RESULTS AND INTERPRETATION The stained smear contains mixture of Gram positive cocci arranged in grape-like clusters (eg. Staphylococcus species) and Gram negative bacilli (eg. E. coli). Various uses of Gram’s staining are summarized in the box 7-2.

Table 7-1 List of Gram positive and Gram negative bacteria

Gram positive bacteria Gram positive cocci In clusters: eg. Staphylococcus aureus In chains: eg., Streptococcus species Lanceolate shaped in pairs: eg. Streptococcus pneumoniae In pairs and short chains: eg., Enterococcus species

Gram positive bacilli In chains eg. Bacillus anthracis with spores eg. Clostridium species

Gram negative bacteria Gram negative cocci In pairs: eg.Neisseria gonorrhoea Neisseria meningitidis

Gram negative bacilli eg. Escherichia coli Klebsiella pneumoniae Comma shaped, curved Gram negative bacilli eg. Vibrio cholerae

Textbook of Practical Microbiology

FIGURE 7-1 Gram stained smear of Staphylococcus aureus, x 1000.

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FIGURE 7-2 Gram stained smear of Gram negative bacilli, x 1000.

BOX: 7-1 MODIFICATIONS OF GRAM’S STAINING 1 Kopeloff and Beerman’s Gram method This method uses acetone as a decolouriser. 2 Jensen’s Gram method for smears This method uses alcohol as decolouriser and weak neutral red as counter stain. This modification is recommended for examination of smears for gonococci and meningococci. 3 Preston and Morrell’s Gram method This method uses iodine-acetone as decolouriser giving good results without the need for careful timing of the decolourisation step.

BOX: 7-2 USES OF GRAM’S STAINING 1 Urine specimens Identifies urine specimens that contain bacteria greater than 105 cfu/ml of urine. Presence of at least 1 organism/ oil immersion field correlates with significant bacteriuria ( > 105 cfu/ml). Presence of polymorphonuclear (PMN’s) in uncentrifuged urine correlates with number of PMN’s excreted per hour. In patients with >400,000 PMN excreted into urine/hour are likely to be infected. 2 Stool specimens Useful to detect Candida. Clostridium difficile. 3 Sputum specimens Useful to detect Streptococcus pneumoniae. 4 Pus specimen Useful to detect Staphylococcus aureus in pus from abscesses. Streptococcus species in exudates from tonsillitis, pharyngitis and impetigo. Gram negative rods in pus from cases of chronic otitis media and post operative wound infections.

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Gram’s Staining

KEY FACTS 1 Gram staining is a differential stain, most commonly employed for diagnostic identification of bacteria in clinical specimens. 2 Gram staining differentiates bacteria into two categories: purple coloured Gram positive and pink coloured Gram negative bacteria. 3 Tissue cells, leucocytes and the debris of inflammatory exudates all stain pink in Gram’s stained smears. 4 Gram positive bacteria have a thicker and denser peptidoglycan layer in the cell walls, which make them less permeable to the stain, in comparison to those of the Gram negative bacteria. 5 The bacterial smear should not be overheated during heat fixation or over decolourised with alcohol, as it can cause in Gram positive bacteria losing the dye-iodine complex and appearing Gram negative. 6 The age of culture can also influence the Gram stain reaction i.e. cultures more than 24 hours old can lose ability to retain dye-iodine complex and appear pink coloured Gram-negative.

VIVA 1 Define differential stain. What are the other differential stains used for staining? Ans. Differential stain is defined as those that contain two different coloured dyes, such that when the staining is completed, different structures will take up different colours and can be differentiated. The other differential stain that can be used for staining is acid fast stain and Albert’s stain. 2 State why Gram’s stain is said to be a differential stain? Ans. Gram’s stain is said to be a differential stain because it differentiates bacteria as Gram positive bacteria (appears violet) and Gram negative bacteria (appears pink). 3. Describe differences between a Gram positive and Gram negative cell wall. Ans. The important differences between Gram positive and Gram negative cell wall are as follows: i) Gram positives have a thick peptidoglycan layer in the cell wall when compared to Gram negatives which possess a thinner peptidoglycan layer. ii) Teichoic acid is present in Gram positive cell wall while it is absent in Gram negative cell wall. iii) Lipopolysaccharide is absent in Gram positive cell wall and present in Gram negative cell wall. 4 Describe the conditions, which may result in a Gram-positive bacteria turning as Gram negative. Ans. The conditions that can result in Gram positive bacteria appearing Gram negative are: i) Excess decolourisation by ethanol/ acetone. ii) Loss of cell wall due to action of lysozyme, penicillin, changes in the pH and aged cultures. 5 Give 2 examples each of Gram positive and Gram negative cocci and bacilli. 6 What are ‘Gram variable’ bacteria? Ans. “Gram variable’ bacteria are those Gram positive bacteria that have lost their cell wall integrity because of antibiotic treatment, old age, or action of autolytic enzymes. These changes allow crystal violet to come out of the cell wall during process of decolourizing resulting in some cells staining pink and others staining purple. 7 Give other examples of decolourizing agents and counter stains. Ans. Other examples of decolourizing agents are acetone, 95% ethanol, acetone- alcohol and iodine-acetone. Other examples of counterstains are basic fuchsin and neutral red.

FURTHER READINGS 1 Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press.) 1996. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins.) 1997. 4 WHO. Guidelines on Standard Operating Procedures for Microbiology. Chapter 4: Staining Techniques. 5 WHO. Manual of Basic Techniques for a Health Laboratory, 1980.

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LESSON

8

27

Acid-Fast Staining

LEARNING OBJECTIVES

stain (methylene blue) appearing blue, while the acid-fast cells retain the red colour of primary stain (carbol fuchsin).

After completing this practical you will be able to : 1 Stain smears by acid-fast staining method. 2 Become familiar with the chemical basis of the acid-fast stain

Different modifications of acid-fast staining is summarized in the box 8-1.

REQUIREMENTS INTRODUCTION The acid fast staining method is a modification of Ehrlich’s (1882) method. Ehrlich suggested the method for the differential staining of tubercle bacilli and other acid-fast bacilli with anilinegentian violet followed by strong nitric acid. The ordinary aniline dye solutions do not readily penetrate the substance of the tubercle bacillus and therefore is unsuitable for staining. The Ziehl-Neelsen acid-fast staining method has proved to be most useful for staining acid fast bacilli belonging to the genus Mycobacterium especially Mycobacterium tuberculosis and Mycobacterium leprae, and also for Nocardia. These acid fast bacilli have a high concentration of the lipid-mycolic acid in their cell walls. When stained they appear pink against a blue background.

PRINCIPLE Acid fastness of acid-fast bacilli is attributed to the presence of large quantities of unsaponifiable wax fraction called mycolic acid in their cell wall and also the intactness of the cell wall. The degree of acid fastness varies in different bacteria. In this staining method, application of heat helps the dye (a powerful staining solution containing carbol fuchsin and phenol) to penetrate the tubercle bacillus. Once stained, the stain cannot be easily removed. The tubercle bacilli resist the decolourizing action of acid-alcohol which confers acid fastness to the bacteria. The other microorganisms, which are easily decolourised by acid-alcohol, are considered non-acid fast. The non-acid fast bacilli readily absorb the colour of the counter

I Equipments Compound light microscope. II Reagents and glass wares Bunsen flame/ torch soaked in methylated spirit, loop wire, glass slides, slide rack, strong carbol fuchsin, acid-alcohol (3 ml HCl + 97 ml ethanol) (decolourising agent), and Loeffler’s methylene blue (counter stain). Preparation of strong carbol fuchsin: This solution is prepared by dissolving 5 grams basic fuchsin powder in 25 grams crystalline phenol by placing them in a 1 litre flask. The flask containing solution is kept over a boiling water-bath for about 5 minutes, shaking the contents from time to time. When the solution is complete, 50 ml of 95% alcohol or 100% ethanol is added to the solution and mixed thoroughly. Then 500ml of distilled water is added to it and the mixture is filtered before use. Preparation of 20% sulphuric acid : 800ml of water is collected in a large flask. The 200ml concentrated sulphuric acid (about 98% or 1.835g / ml)) is poured slowly down the side of the flask into the water, about 50 ml at a time. The mixture becomes hot. Remainder of acid is added in same manner. Note: The acid must be added to the water. It is dangerous to add the water to the acid. Great care must be taken to avoid spilling the acid on skin, clothing or elsewhere. Preparation of 95% alcohol : This is prepared by adding 95 ml of ethanol and adding water to it to make 100ml.

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Acid Fast Staining

Preparation of acid-alcohol decolouriser: This solution contains 75 ml concentrated hydrochloric acid (HCl) and 25 ml of industrial methylated spirit. Methylated spirit is poured into a large flask. The flask is placed in 5–8 cmm of cold water in the sink. Then hydrochloric acid is added slowly and the top of the flask is covered to stop the fumes from escaping. It is left for 10 minutes. It is then decanted into a labeled bottle for use. The final concentration of HCl is 3%. III Specimen Sputum smear positive for tubercle bacilli / culture smear of Mycobacterium species. Note: Frequently examined specimens for the detection of Mycobacterium tuberculosis are summarized in the box 8-2.

PROCEDURE 1 Heat fixes the smears by passing the slide 2–3 times gently over the flame with the smear side up. Allow the smear to be air dried. 2 Put the smears on a slide rack and cover the smears with strong carbol fuchsin. Allow it to stain for 5 minutes. 3 During this period, heat the slides from below intermittently by Bunsen flame or torch soaked in methylated spirit without boiling the solution, until the steam rises. Do not allow the stain to dry on the slide, and if necessary add more carbol fuchsin to cover the smear. 4 Rinse the smears gently under tap water. 5 Cover the smear with 20% sulphuric acid for at least 10 minutes for decolourisation. 6 Wash the slides thoroughly with water to remove all traces of acid. Note: Decolourisation with 95% alcohol for 2 minutes is only optional and may be omitted. 7 Cover the smear with Loeffler’s methylene blue for 15–20 seconds.

FIGURE 8-1 Acid fast stained smear of Mycobacterium tuberculosis, x 1000.

8 Rinse the smears again under tap water and air dry it. 9 Observe the smear first under low power (10x) objective, and then under oil immersion (100x) objective. Note: The smear should be examined following a zig-zag pattern for at least 10 minutes or 300 fields, before declaring the smear negative. 10 Record the observations in the note book. Findings are recorded, together with grading of the positive smear.

QUALITY CONTROL A positive control sputum smear from a known case of tuberculosis patient, stained with Z-N stain is compared with the stained test smear for appropriate morphology and staining appearance. With appropriate staining, the acid-fast bacillus appears pink against blue background of pus cells.

OBSERVATION Presence of 3 x 0.3 µm sized pink coloured slender rod shaped structures are seen with curved ends, and are scattered amidst blue coloured round cells with darkly stained multilobed nucleus.

RESULTS AND INTERPRETATION The stained smear contains pink coloured acid fast bacilli seen among the blue coloured multilobed pus cells. The smear is positive for acid fast bacilli. Probably, the smear contains Mycobacterium tuberculosis (Fig. 8-1). Note: List of other acid–fast structures are provided in the table 8-1.

FIGURE 8-2 Acid fast stained smear of Mycobacterium leprae, x 1000

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Table 8-1 List of acid –fast structures Bacteria

Parasites

Fungi

Other structures

Mycobacterium tuberculosis ( Fig. 8-1) Mycobacterium leprae ( Fig. 8-2) Mycobacterium smegmatis Mycobacterium fortuitum Nocardia asteroids Nocardia brasiliensis Nocardia caviae

Cryptosporidium parvum Cyclospora cayetanensis Isospora belli

Fungal spores

Spermatozoa head

BOX 8-1 DIFFERENT MODIFICATIONS OF ACID FAST STAIN AND THEIR USES 1 5% sulphuric acid is used as a decolourizing agent for staining Mycobacterium leprae. 2 1% sulphuric acid is used as a decolourizing agent for staining Nocardia species, Cryptosporidium and Isospora oocysts (Kinyoun’s modification of acid-fast stain). 3 0.25% sulphuric acid is used as a decolourizing agent for staining spores.

BOX 8-2 FREQUENTLY EXAMINED SPECIMENS FOR THE DETECTION OF MYCOBACTERIUM TUBERCULOSIS Pulmonary tuberculosis Sputum. Bronchial or laryngeal washings. Gastric lavage (when sputum is swallowed as in children). Miliary tuberculosis Bone marrow. Liver biopsy. Tuberculous meningitis Cerebrospinal fluid. Renal tuberculosis Urine.

KEY FACTS 1 2 3 4

Ziehl-Neelsen stain is intended for the differential staining of tubercle bacilli from other acid fast bacilli. Acid fast bacilli appear pink coloured in stained smears. At the end of the process of decolourisation by sulphuric acid, the smear will appear faintly pink. Acid fastness of the bacteria is attributed to presence of mycolic acid in high concentration in the cell walls of tubercle bacilli and also to the intactness of the cell wall. 5 Positive sputum smear is graded only under oil-immersion as follows: 3–9 AFB in entire smear = 1+ 10 or > AFB in entire smear = 2+ 10 or > AFB in each oil immersion field = 3+

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Acid Fast Staining

VIVA 1. List the acid fast organisms. 2. Why are some bacteria acid fast? Ans. Some bacteria are acid-fast because their thick waxy cell wall is made up of long chain fatty (mycolic) acids. These mycolic acids render the cells resistant to decolourisation, even with acid-alcohol decolourisers. Thus when acid is added as a decolouriser, these acid fast bacteria retain the dye because the dye is more soluble in the cytoplasm compared to sulphuric acid. 3 How do you grade a positive sputum smear? 4. List another method that can be used for detecting acid fast bacteria in a smear? Ans. Fluorescent staining methods using fluorochrome dyes such as auramine O and rhodamine is another method that can be used for detecting acid fast bacteria in a smear. By this method acid fast bacteria fluoresces bright yellow or orange against a greenish background. 5. What are the various specimens obtained in the laboratory for the diagnosis of tuberculosis? 6. List different modifications of acid fast staining and their uses.

FURTHER READINGS 1 Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press.) 1996. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins.) 1997. 4 WHO. Guidelines on Standard Operating Procedures for Microbiology. Chapter 4: Staining Techniques. 5 WHO. Manual of Basic Techniques for a Health Laboratory, 1980.

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LESSON

9

31

Albert’s Staining

LEARNING OBJECTIVES

II Reagents and glass wares Bunsen flame, loop wire, glass slides, Albert’s stain I and II.

After completing this practical you will be able to: 1 Stain smears by Albert’s staining to demonstrate the presence of metachromatic granules in the diphtheria bacillus.

INTRODUCTION The diphtheria bacillus, Corynebacterium diphtheriae has well developed granules within their bacterial cytoplasm. These granules are known as Volutin granules, Babes Ernst granules, polar bodies or metachromatic granules. These granules are made up of polymetaphosphate and are seen in unstained wet preparations as round, refractile bodies within the bacterial cytoplasm. With basic dyes, granules tend to stain more strongly than the rest of the bacterium. With toluidine blue or methylene blue, they stain metachromatically, and appear reddish purple in colour. These granules are clearly demonstrated best by special stains such as Albert’s, Neisser’s or Puch’s stain. With Albert’s staining, the bacilli appear green with bluish-black metachromatic granules.

Preparation of Albert’s stain I: This stain is composed of 1.5 grams toluidine blue, 2 grams malachite green, 10 ml glacial acetic acid, 10 ml alcohol (95% ethanol) and 1 litre distilled water. Toluidine blue and malachite green are dissolved in the alcohol and then added to the water and acetic acid. The stain is then allowed to stand for one day and then filtered. Note: Toluidine blue stains the granules bluish black due to metachromatic effect and malachite green stains the bacilli green. Preparation of Albert’s stain II: Albert’s II (also known as Albert’s iodine) is composed of 6 gram iodine, 9 gram potassium iodide and 900 ml distilled water. The solution is made by first dissolving 2 gram potassium iodide in 10 ml distilled water and then 1 gram of iodine is further added to it with dissolution. Then 290 ml of distilled water is added and final volume is made up to 300 ml. III Specimen Exudate smear collected directly from pseudomembrane obtained using a throat swab / culture smear of C. diphtheriae.

PRINCIPLE The granules present in the diphtheria bacilli exhibit metachromasia property and hence appear bluish-black coloured when stained with the toluidine blue present in Albert I reagent, while the diphtheria bacillus appears green due to malachite green present in Albert I reagent.

REQUIREMENTS I Equipments Compound light microscope.

PROCEDURE 1 Heat fix the smears by passing the slide 2–3 times gently over the flame with the smear side up. Allow the smears to be air dried. 2 Put the smears on a slide rack and cover the smears with Albert’s stain I. Allow it to stain for 3-5 minutes. 3 Rinse the smears gently under tap water and blot those dry. 4 Then cover the smear with Albert’s stain II. Allow it to act for 1 minute. 5 Rinse the smears again under tap water and blot those dry. 6 Observe the smear first under low power (10x) objective, and then under oil immersion (100x) objective.

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Albert’s Staining

7 Record the observations in the note book. Findings are recorded, together with grading of the positive smear.

The smear is positive for bacilli showing bluish-black metachromatic granules. Probably, the smear contains Corynebacterium diphtheriae.

QUALITY CONTROL A known positive control smear of the diphtheria bacillus stained with Albert’s stain and the stained test smear are compared for appropriate morphology and staining appearance. With appropriate staining, the bacillus should appear green with bluish black metachromatic granules.

OBSERVATION 1–2 µm sized green coloured bacilli showing Chinese letter arrangement at angles to each other, containing bluish-black metachromatic granules, are seen (Fig. 9-1).

RESULTS AND INTERPRETATION

FIGURE 9-1 Albert’s stained smear of Corynebacterium diphtheriae showing volutin granules, x 1000.

The stained smear contains malachite green stained bacilli showing bluish-black metachromatic granules.

BOX 9-1 RAPID STAINING BY DIRECT FLUORESCENT ANTIBODY METHOD Direct fluorescent antibody testing for Corynebacterium diphtheriae is a rapid diagnostic method like that of Albert’s staining. This method involves the use of specific antibodies raised against C. diphtheriae conjugated with a fluorescent dye to detect directly the C. diphtheriae antigen in the throat swab or smear taken from pseudomembrane. The smear is viewed by immunofluorescent microscopy. The method is highly specific as well as sensitive.

VIVA 1. What is metachromasia? Ans. When a substance is stained with a particular coloured dye, and if a change in original colour is observed, this phenomenon is called metachromasia. This phenomenon is observed with C. diphtheriae. The granules present in the bacteria are called metachromatic granules because the blue colour of the stain is changed to bluish-black by those granules. 2 Mention the other names for metachromatic granules. 3 What is the composition of metachromatic granules? Ans. Metachromatic granules are composed of polymetaphosphate, ribonucleic acid and protein. 4 What is the significance of metachromatic granules? Ans. The significance of metachromatic granules is that they represent storage depots of materials needed to form highenergy phosphate bonds. They are not found during active growth period and are depleted under starvation conditions. Their presence in thin slender bacilli helps to distinguish C. diphtheriae from short, thick, plumpy, non- pathogenic diphtheroids which lack them. 5 What is the typical arrangement of C. diphtheriae? What is the reason for such an arrangement? Ans. The typical arrangement of C. diphtheriae is Chinese letter pattern or Cuneiform arrangement, (bacilli are seen arranged at various angles to each other resembling the letters V or L. This is due to incomplete separation of the daughter cells after longitudinal binary fission. 6 Mention the other stains used for the detection of C. diphtheriae.

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KEY FACTS 1 Volutin granules of C. diphtheriae are made up of polymetaphosphate. 2 Special stains like Albert’s stain, Neisser’s stain and Puch’s stain are used to demonstrate metachromatic granules. 3 Granules exhibit metachromasia and are seen in unstained wet preparations as round refractile bodies within bacterial cytoplasm. 4 The diphtheria bacillus gives its characteristic volutin staining reactions best in a young culture (18–24 hours) on Loeffler’s serum slope or serum medium.

FURTHER READINGS

1 Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press.) 1996. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins.) 1997. 4 WHO. Guidelines on Standard Operating Procedures for Microbiology. Chapter 4: Staining Techniques. 5 WHO. Manual of Basic Techniques for a Health Laboratory, 1980.

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LESSON

10

Capsule Staining

LEARNING OBJECTIVES After completing this practical you will be able to : 1 Demonstrate capsule of the bacteria present in animal tissues, blood, serous fluids and pus and artificial cultures , by positive staining and negative staining methods. 2 Differentiate between capsulated and non-capsulated bacteria.

the capsular materials are water soluble and may be dislodged and removed with vigorous washing. Bacterial smears should not be heated, because the resultant cell shrinkage may create a clear zone around the organism, an artifact that can be mistaken for capsule. The capsule is non-ionic, so that the dyes commonly used will not bind to it. Two dyes, one acidic and one basic, are used to stain the background and the cell wall, respectively.

REQUIREMENTS INTRODUCTION Some bacteria secrete chemical substances that accumulate on the outer surfaces of the cell wall and form capsules. When the capsule has a diameter of 20 nanometer or more and seen under light microscope, it is referred to as macrocapsule. When the diameter is less than 20 nm and seen under electron microscope, it is called microcapsule. Capsules may be seen in stained or unstained preparations as a clear zone around the bacteria. Two types of staining procedures can be employed to demonstrate the capsule. They are the positive and negative staining procedures. In the positive staining technique, the capsule is stained and coloured whereas in the negative staining procedure, the background is stained and the capsule is seen as unstained hallow around the organism. The best method for staining capsules on bacteria in either liquid or solid media is the wet-film India ink method. Dry-film negative staining methods using India ink, nigrosin or eosin are somewhat less reliable, since occasionally shrinkage spaces give the appearance of capsules around bacteria that are noncapsulated.

PRINCIPLE Chemically, the capsular material is a polysaccharide, a glycoprotein or a polypeptide. Capsule staining is more difficult than other types of differential staining procedures because

I Equipments Compound light microscope. II Reagents and glass wares Bunsen flame, inoculating loop, staining tray, glass slides, 1% crystal violet and 20% copper sulphate (CuSO4 5H2O) for positive staining; and India ink or Nigrosin stains for negative staining. Nigrosin staining is prepared by adding 0.03 grams of nigrosin in 100 ml of distilled water. III Specimen Culture of Streptococcus pneumoniae (A capsulated bacterium).

PROCEDURE For positive staining of smears 1 Make a smear from colony of S. pneumoniae on a clean grease free glass slide, and allow it to air dry. Note: The smear is not heat fixed. 2 Put the smear on a slide rack and flood smear with crystal violet. Allow it to stain for 5-7 minutes. 3 Wash the smear with 20% copper sulphate solution and blot it dry. 4 Observe the smear first under low power (10x) objective, and then under oil immersion (100x) objective. 5 Record the observations in the note book.

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For negative staining of smears 1 Take a clean grease free glass slide. 2 Put a large loopful of undiluted India ink on the slide. 3 Then add a small loopful of liquid bacterial culture to the India ink and emulsify. 4 Take a clean, grease free cover slip and place on the ink drop and press it down, so that the film becomes very thin and thus pale in colour. 5 Observe the wet film under high power (40x) objective. 6 Record the observations in the note book.

QUALITY CONTROL The test smear subjected to capsular staining (e.g. both positive and negative staining technique) is compared with known stained control smear of S. pneumoniae (capsulated bacteria) for appropriate structural details and staining appearance.

FIGURE 10-1 India ink staining of Streptococcus pneumoniae showing capsule, x 1000.

OBSERVATION Observation of positive staining method In the culture smear, the capsule is seen as a light blue hue in contrast to the deep purple colour of the cell. Observation of negative staining method The capsule in negative staining method is seen as clear refractile, halo around the organism against a black background.

RESULTS AND INTERPRETATION Positive staining method: The smear shows capsulated bacteria. e.g. S. pneumoniae. Negative staining method: The wet India-ink film contains capsulated bacteria. e.g. S. pneumoniae (Fig. 10-1).

FIGURE 10-2 Gram’s staining showing clear unstained capsulated area around Streptococcus pneumoniae, x 1000.

KEY FACTS 1 Capsule of the bacteria, Bacillus anthracis in the stained blood smears is demonstrated by McFadyean reaction which uses polychrome methylene blue. 2 Capsules of the bacteria can be demonstrated by two methods: positive staining and negative staining procedures. 3 In the positive staining technique, the capsule is stained and coloured. In negative staining, the background is stained, and the capsule is seen as an unstained halo around the organism. 4 The wet India ink preparation can also be employed for demonstration of slime which are irregular masses of amorphous material seen lying between the bacteria and outside the capsules of capsulate ones. 5 Modified India ink preparation using 2% mercurochrome helps clearly demonstrate the internal structure of capsulated budding yeast. 6 India ink film should have appropriate thickness if the film is too thick, the capsule will be obscured by overlying ink and if too thin, the capsules get crushed or flattered and the India ink background becomes too pale to give a good contrast.

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Capsule Staining

VIVA 1 Give examples of bacteria having polysaccharide or polypeptide capsules. Ans : a) Polysacchride capsule Staphylococcus aureus Group B streptococci Group D streptococci Streptococcus pneumoniae Neisseria meningitidis Haemophilus influenzae Anaerobic Gram negative bacilli b) Polypeptide capsule Bacillus anthracis 2 List various capsulated bacteria Staphylococcus aureus (microscopically visible capsules) Streptococcus pyogenes (some stains) Group C Streptococci ( capsules made of hyaluronic acid). Group B streptococci Group D streptococci (polysacchaide capsules) Streptococcus pneumoniae. Neisseria memigitidis (fresh isolates capsulated) Bacillus anthracis (polypeptide capsule) Clostridium perfringens Bacteriodes fragilis Escherichia coli (some strains) Klebsiella pneumoniae Vibrio parahaemolyticus Yersinia pestis Francisella tularensis Haemophilus influenzae Bordetella pertusis 3 Describe the functions of the capsule. Ans: i) Capsules contribute to the virulence of pathogenic bacteria by inhibiting phagocytosis, ii) They protect the bacteria from antibody, and iii) Capsular material is antigenic in nature and may be demonstrated by serological methods such as Quellung reaction widely employed for typing of pneumococci. 4 Describe the capsular antigens Ans: i) Type specific capsular polysaccharide antigen of pneumococcus is also known as specific soluble substance. ii) Capsular polysaccharide antigens of meningococci have been used to classify them into 13 serogroups, and iii) K antigen (acidic capsular polysaccharide antigen) of Escherichia coli protect it from bactericidal effect of complement and phagocytes.

FURTHER READINGS 1 Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press.) 1996. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins.) 1997. 4 WHO. Guidelines on Standard Operating Procedures for Microbiology. Chapter 4: Staining Techniques. 5 WHO. Manual of Basic Techniques for a Health Laboratory, 1980.

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LESSON

11

Spore Staining

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Stain smears for bacterial endospores by malachite green stain. 2 Differentiate between bacterial spore and vegetative cell forms in the bacterial smear.

INTRODUCTION Spore production is a very important characteristic of some bacteria such as members of anaerobic genera Clostridium and aerobic genus Bacillus. They are highly resistant and metabolically inactive forms. They occur when environmental conditions become unfavorable for continuing vegetative cellular activities, particularly with the exhaustion of nutritional carbon source. Because of the chemical composition of spore layers, the spore is resistant to deleterious effects of excessive heat, freezing, radiation, desiccation, and chemical agents, as well as to the commonly employed microbiological stains. The morphology of the bacterial endospores is best observed in unstained wet films under the phase-contrast microscope, where they appear freely or within the bacterial cells as large, refractile, oval or spherical bodies. Different staining techniques are available for staining of spores. These are malachite green stain (Schaeffer and Fulton method), modified Ziehl-Neelsen stain using 0.25% sulphuric acid as decolouriser, toluidine blue stain and also Gram’s stain. With the Gram’s stain, the body of the bacillus appears deeply stained, whereas the spore is unstained and appears as a clear area in the organism. The spores on staining with modified Ziehl-Neelsen stain appear red and bacilli blue.

PRINCIPLE Malachite green stain, also known as Schaeffer-Fulton Method for bacterial endospores uses two different reagents: primary

stain (malachite green) and counter stain (0.5% safranine or 0.05% basic fuchsin).Ordinary tap water acts as decolourising agent. Unlike most of the vegetative cells that are stained by common procedures, the spore, because of its impervious coats, are not stained by the primary stain easily. The application of heat facilitates penetration of the primary stain, malachite green. After the primary stain is applied and the smear is heated, both the vegetative cell and spore appear green. Once the spore is stained with the malachite green, it cannot be decolourised by tap water, which removes only the excess primary stain. The spore remains green. On the other hand, water removes the stain from the vegetable cells, because the stain does not demonstrate a strong affinity for the vegetative cell components and these vegetable cells therefore become colourless. Red coloured-safranine as counterstain is used as the second reagent to colour the decolourised vegetative cells, which will absorb the counterstain and appear red. The spores retain the green of the primary stain.

REQUIREMENTS I Equipments Compound light microscope. II Reagents and lab wares Bunsen burner, beaker of boiling water, staining tray, glass slides, inoculating loop, malachite green and safranine. Preparation of malachite green stain: This stain is prepared by dissolving 5 gram of malachite green in 100 ml of distilled water. Preparation of safranine stain: This stain is prepared by dissolving 0.5 gram of safranine in 100 ml distilled water.

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Spore Staining

III Specimen Smear collected from 48 hours to 72 hours nutrient agar slant culture of Bacillus cereus/ thioglycollate culture of Clostridium butyricum. On a clean glass slide, a smear from the culture is made in saline, then air dried and fixed with heat.

Bacterial endospores stain green, and vegetative bacilli stain red. (Fig. 11-1)

PROCEDURE

RESULTS AND INTERPRETATION

1 Heat fix the smears by passing the slide 2–3 times gently over the flame with the smear side up. Allow the smear to be air dried. 2 Put the slide with the smear over a beaker of boiling water, resting it on the run with the bacterial film upper most. 3 When, within several seconds, large droplets have condensed on the underside of the slide, flood the smear with 5% acqueous solution of malachite green and allow to act for 1 minute, while the water continues to boil. 3 Wash the smears with cold water. 4 Then cover the smear with 0.5% safranine or 0.05% basic fuchsin. Allow it to act for 30 seconds. 5 Rinse the smears again under tap water and blot those dry. 6 Observe the smear first under low power (10x) objective , and then under oil immersion (100x) objective. 7. Record the observations in the note book.

A 2 – 3 µm red coloured rod-shaped structure seen along with an intracellular 0.5 µm sized spherical green coloured structure. It represents red coloured vegetative bacilli with green coloured spores by the malachite green staining method. May be sporebearing bacilli (eg. Bacillus species or Clostridium species)

OBSERVATION

QUALITY CONTROL A check smear of a known positive control of bacteria with spores is stained with the malachite green staining method, and its appropriately stained morphology is observed under the microscope and compared with the stained test smear.

FIGURE 11-1 Spore staining, x 1000.

VIVA 1 Name spore bearing bacilli causing anthrax and gas gangrene. Ans : Spore bearing bacilli causing anthrax is Bacillus anthracis. Spore bearing bacilli causing gas gangrene are Clostridium perfringens , Clostridium novyi , Clostridium septicum , and Clostridium histolyticum. 2 Spores of which .bacilli are used to test efficacy of sterilization using i) dry heat and ii) moist heat? Ans: The spores of non toxigenic strain of Clostridium tetani are used for testing efficacy of sterilization by dry heat. The spores of Bacillus stearothermophilus are used for testing efficacy of sterilization by moist heat. 3 Which sterilization method is employed for destruction of spores? Ans: Autoclaving at 120°C for 15 minutes is the sterilization method employed for destruction of spores in clinical specimens.

KEY FACTS 1 Spores are highly resistant, metabolically inactive forms occurring when environmental conditions are unfavorable. 2 The morphology of spores is best observed in unstained wet films under the phase-contrast microscope. 3 The staining methods available for demonstrating spores are Gram’s staining, modified acid fast staining using 0.25% sulphuric acid as decolouriser, toluidine blue staining and malachite green staining (Schaeffer-Fulton Method). 4 With malachite green staining vegetative bacilli appear red and spores green. 5 With toluidine blue staining , vegetative bacilli appear light blue and spores dark blue.

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FURTHER READINGS 1 Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press.) 1996. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins.) 1997. 4 WHO. Guidelines on Standard Operating Procedures for Microbiology. Chapter 4: Staining Techniques. 5 WHO. Manual of Basic Techniques for a Health Laboratory, 1980.

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LESSON

12

Negative Staining

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to:

I Equipment Compound light microscope.

1 Perform a negative staining procedure for demonstration of bacteria

INTRODUCTION Negative staining procedure is so called because the background gets stained and the organism remains colourless. It is also known as ‘Indirect staining. The procedure requires the use of acidic stains such as India ink or Nigrosin. Negative staining finds its utility for the demonstration of capsule and bacteria difficult to stain such as Treponema palladium. Wet film India-ink method is the best method for staining capsules of bacteria from cultures in either liquid or solid media. A modification of India ink using 2% mercurochrome helps to clearly demonstrate the capsulated budding yeast, Cryptococcus neoformans. Indian ink method for demonstration of capsule is described earlier in the chapter 10. In this chapter dry film negative staining by using nigrosin to detect bacteria in dry smears will be described.

PRINCIPLE The acidic dyes such as India ink, nigrosin or eosin have negatively charged chromogen, and will not readily combine with the negatively charged bacterial cytoplasm. Instead it forms a deposit around the organism, leaving the organism itself colourless. Therefore, the unstained cells are easily discernible against the coloured background.

II Reagents and glass wares These include Bunsen flame, staining tray, glass slides and coverslips nigrosin stains. Nigrosin staining solution is prepared by adding 0.03 gram of nigrosin in 100 ml of distilled water. III Specimen 24 hour broth culture of Klebsiella pneumoniae (A capsulated bacterium).

PROCEDURE 1 Take a clean grease free glass slide. 2 Put a small drop of nigrosin close to one end of a clean slide. 3 Using a sterile loop, a loopful of broth culture of the capsulated organism is mixed with the nigrosin drop. 4 With the edge of a second slide, held at 30° angle and held in front of the bacterial suspension mixture, spread the drop along the edge of the applied slide. The slide is then pushed away from the previously spread drop of suspended organism, forming a thin smear. 5 Air dry the preparation without any heat fixation. 6 Observe the stained smear under oil immersion (100x) objective. 7 Record the observations in the note book.

QUALITY CONTROL The nigrosin-stained preparation of test organism is compared with the preparation of capsulated control organism such as Streptococcus pneumoniae, for appropriate staining and capsule demonstration.

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OBSERVATION The capsule or the bacterial organism is seen as a clear halo against a black or dark background in the wet film or dry film preparation (Fig. 12-1).

RESULTS AND INTERPRETATION The nigrosin-stained smear contains capsulated bacteria. e.g. K. pneumoniae.

FIGURE 12-1 Nigrosin-stained smear showing capsulated K. pneumoniae, x 400.

BOX 12-1 LIST OF MOST COMMON CAPSULATED ORGANISMS THAT CAN BE DEMONSTRATED BY NEGATIVE STAINING Bacteria Streptococcus pneumoniae Klebsiella pneumoniae Haemophilus influenzae Group B streptococci Group D streptococci

Fungus Cryptococcus neoformans

KEY FACTS 1 Negative staining procedure is so called because the background gets stained and the organism remains colourless. 2 In negative staining method only background is stained, the organism and capsule remained unstained and refractile.

VIVA 1 List the different stains that can be employed for negative staining. Ans: The different stains employed for negative staining are: India ink, modified India ink (using 2% mercurochrome) and 10% nigrosin. 2 List the capsulated organisms and bacteria that can be demonstrated by negative staining. 3 What is negative staining? Which bacteria can be demonstrated by India Ink. Ans: In negative staining, the bacteria are mixed with dyes such as India ink or nigrosin that provide a uniformly coloured background against which the unstained bacteria stand out in contrast. This is particularly useful in the demonstration of bacterial capsules.

FURTHER READINGS 1 Evans EGV, Killington RA, Heritage J. Introductory Microbiology. (Cambridge University Press.) 1996. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins.) 1997. 4 WHO. Guidelines on Standard Operating Procedures for Microbiology. Chapter 4: Staining Techniques. 5 WHO. Manual of Basic Techniques for a Health Laboratory, 1980.

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UNIT

III Cultivation of Bacteria

Lesson 13

Media for Routine Cultivation of Bacteria

Lesson 14

Temperature Requirement for Growth of Bacteria

Lesson 15

pH Requirement for Growth of Bacteria

Lesson 16

Oxygen Requirement for Growth of Bacteria

Lesson 17

Culture of Anaerobic Bacteria

Lesson 18

Sterilization of Commonly Used Culture Media

Lesson 19

Antiseptics and Disinfectants

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LESSON

13

Media for Routine Cultivation of Bacteria

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know the following commonly used culture media as well as their uses in a clinical microbiology laboratory: a. b. c. d. e.

Basal medium Differential medium Selective medium Enriched, and Enrichment media

INTRODUCTION Cultivation of bacteria is often the first step in the diagnosis of infectious disease. Since bacteria have varied growth requirements, a wide range of media is available. The choice of most appropriate media depends on many factors including nutritional and growth requirements of the bacteria, biochemical properties and many other properties of the bacteria.

2 Enriched medium: These are solid selective media. These media, in addition to basal nutrients also contain nutritional supplements like blood, serum, etc., which favour the growth of fastidious bacteria. e.g. blood agar (Fig. 13-2), chocolate agar (Fig. 13-3) , Löwenstein- Jensen medium (Fig. 13-4), etc. 3 Enrichment media: These are liquid selective media. They favor the growth of some bacteria by extending the lag phase of others eg. Selenite F broth. 4 Selective media: These media contain ingredients that selectively enable the growth of some species, while inhibiting others eg. Deoxycholate citrate agar (DCA) medium. This medium is a selective medium for growth of Salmonella spp. present in stool which contains a mixed bacteria flora. This medium inhibits Escherichia coli and other Gramnegative bacteria. 5 Differential media: These media differentiates between species of bacteria depending on a specific property. Example: MacConkey (Fig. 13-5) agar is a differential medium. This medium is used to demonstrate lactose fermenting properties, and differentiate between lactose and non-lactose fermenting bacteria.

PRINCIPLE A number of media have been formulated for growing bacteria (Table 13-1). Media generally contain a carbon source, nitrogen source and some essential minerals and salts. Some media may contain additional nutritional supplements. In addition solid media contain agar as a solidifying agent. Meat extract and peptone are the commonest sources of carbohydrates and amino acids. Media are of different types. These are: 1 Basal media: These contain nutrients that support the growth of non-fastidious bacteria. They do not confer any selective advantage, e.g. nutrient agar (Fig. 13-1).

FIGURE 13-1 Nutrient agar.

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QUALITY CONTROL 1 One un inoculated set of media as sterility control 2 Nutrient agar: Colonies of non-fastidious bacteria such as S. aureus. 3 Blood agar: Haemolytic strain of S. aureus streaked on the plate surrounded by a zone of hemolysis. 4 MacConkey agar: Pink, lactose fermenting colonies of E. coli and colorless colonies of Proteus spp. 5 Selenite F broth: Growth positive Salmonella spp, and growth negative Proteus spp. FIGURE 13-2 Blood agar.

OBSERVATIONS All the inoculated bacteria (e.g. S. aureus, E. coli, P. mirabilis and Salmonella spp) produce colonies on the nutrient agar (basal medium) and blood agar (enriched medium). In addition S. aureus may or may not produce haemolysis on the blood agar.

FIGURE 13-3 Chocolate agar.

REQUIREMENTS I Equipments Bacteriological incubator.

FIGURE 13-4 Lowenstein-Jenson’s medium.

II Reagents and media Different kinds of media such as nutrient agar, blood agar, MacConkey agar and Selenite F broth. III Specimen 24 hour broth cultures of Staphylococcus aureus, E. coli, Proteus mirabilis and Salmonella spp.

PROCEDURE 1 Inoculate a loopful of the test organism, using a sterile inoculating loop, into appropriately labeled plates and tubes. 2 Incubate the plates and tubes for 18 hours at 37°C. 3 Examine the plate and tubes for growth and record observations.

FIGURE 13-5 Mc Conkey agar.

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Media for Routine Cultivation of Bacteria

On MacConkey agar E. coli produces pink, lactose fermenting colonies where as Proteus spp. produces colorless non-lactose fermenting colonies. In SF broth most bacteria are inhibited. Selenite F broth inhibits all except S. Typhi

RESULTS AND INTERPRETATION Different colonies of various bacteria on different media show different growth requirement of these bacteria.

Table 13-1 List of different type of media commonly used for isolation of bacteria Type of medium

Example

Bacteria grown/isolated

Basal media

Nutrient agar Peptone water Blood agar Brain heart infusion agar Alkaline peptone water Selenite F Broth Pikes medium Cetrimide agar MacConkey agar, CLED medium

S.aureus, E.coli, Klebsiella, Proteus, Pseudomonas S.aureus, E.coli, Klebsiella, Proteus, Pseudomonas. S. pyogenes. H. influenzae. V. cholerae. Salmonella spp. Streptococcus spp. Pseudomonas spp. E.coli, Proteus spp., S.aureus, Klebsiella. Proteus, Pseudomonas, Enterobacter, Citrobacter.

Enriched media Enrichment media Selective media Differential media

KEY FACTS 1 Bacteria have different nutritional requirements. Most of the bacteria grow on enriched media. 2 S. aureus can grow on MacConkey medium. 3 Media serve many purposes such as culture and isolation of the bacteria from clinical specimens, demonstration of growth and biochemical characteristics of bacteria, and for storage of bacterial isolates, etc.

VIVA 1 Give examples of different kinds of media used for culture of bacteria. 2 What are the constituents of MacConkey agar? Ans. These are the peptone (2%), sodium taurocholate (0.5%), lactose (1%), neutral red (0.7%), and agar (2%). 3 Name the enrichment medium used for Vibrio cholerae. 4 Mention an important difference between enriched medium and enrichment medium. Ans Enriched medium (e.g., blood agar) is a solid medium where as enrichment medium (e.g., alkaline peptone water) is a liquid medium.

FURTHER READINGS

1 Bhattacharya S, Vijayalakshmi N, Parija SC. Uncultivable bacteria: Implications and recent trends towards identification. Indian J Med Microbiol. 2002: 20; 174-177. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 3 Isenberg HD. (Ed) Clinical Microbiology Procedures Handbook. American Society for Microbiology, Washington DC, 1992. 4 PHLS Standard Operating Procedures. Inoculation of Culture Media. No B.SOP 54 Version: 1, 1998. 5 WHO. Guidelines on standard operating procedures for Microbiology. Blood safety and clinical technology. Chapter 6: Cultivation of bacteria on laboratory media. (World Health Organisation, Geneva) 1997.

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LESSON

14

Temperature Requirement for Growth of Bacteria

LEARNING OBJECTIVES After completing this practical you will be able to understand: 1 Effect of temperature on growth of the bacteria in culture media.

INTRODUCTION A number of factors influence the growth and culture of bacteria on culture media. Apart from nutritional requirements, which vary greatly, environmental factors also play an important role. These factors include availability of proper temperature, pH and oxygen in the culture environment.

PRINCIPLE Cellular enzymes of the bacteria are optimally active at a certain temperature. Above this point, they get denatured. On the other hand, at low temperatures, they tend to get inactivated. Either event causes biochemical reaction in bacterial cells to cease, thereby interfering with the survival of the organism. In general, bacteria can survive across a wide range of temperatures from 5 0° C to 80°C. However, each bacterial species has an optimum growth temperature at which reproduction is most rapid and also a maximum/minimum growth temperature beyond which growth does not occur. The ideal temperature for growth may not coincide with that for specific enzyme activities. Based on their temperature requirements, bacteria can be classified into the following groups: 1 Psychrophilic bacteria: These bacteria can grow within a temperature range of 5°C to 20°C. Optimum temperature for growth of these bacteria varies between 10 °C to 15°C. 2 Mesophilic bacteria: These bacteria can grow within a temperature range of 20°C to 45°C. Optimum temperature for growth of these bacteria is 37°C (human body temp). These

bacteria prefer to grow in the bodies of warm blooded hosts and include most bacterial pathogens that cause diseases in humans. 3 Thermophilic bacteria: These bacteria can grow above 35°C. They are either facultative with an optimum temperature of 45°C -60°C but capable of growth at 37°C or obligate, growing only at temperature above 50°C. Examples of bacteria showing different temperatures for their growth is summarized in the table 14-1.

REQUIREMENTS I Equipments Bacteriological incubator. II Reagents and media Nutrient agar plates. III Specimen A 24 hour broth culture of Serratia marcescens, Escherichia coli, Enterococcus faecalis and Flavobacterium spp

PROCEDURE 1 Mark the nutrient agar plates into 4 quadrants each. 2 Using a sterile loop, aseptically inoculate a drop of each culture, taking care to label each quadrant correctly. 3 Incubate the plates at 4°C, 20°C, 37°C and 60°C for 24-48 hr. 4 Observe the plates for growth of bacteria and pigment production if any.

QUALITY CONTROL 1 Nutrient agar plate inoculated with Proteus mirabilis as quality control for mesophilic bacteria.

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2 Nutrient agar plate inoculated with Pseudomonas aeruginosa as quality control for psychrophilic bacteria. 3 Nutrient agar plate inoculated with Bacillus stearothermophilus as quality control for thermophilic bacteria.

OBSERVATIONS 1 Reproduction of bacteria is maximal at optimal temperature and will be seen as a luxuriant growth of colonies. 2 Pigment production is often optimum at a temperature different from that for growth. S. marcescens produces a red/magenta pigment.

RESULTS AND INTERPRETATION 1 Growth of all species tested will be absent at 4°C except for Yersinia enterocolitica and Flavobacterium spp. 2 At 20°C growth will be minimal/inhibited for E. coli, S. marcescens and Flavobacterium spp. Flavobacterium spp is capable of growth at low temperature as well. 3 At 37°C all the bacterial species tested will produce good growth. Pigment production will be seen with the cultures of both Flavobacterium and S. marcescens. Pigment production is influenced by temperature with a greater intensity on color at 20°C for both the species tested. There is a different temperature requirement for optimal growth and pigment production. 4 At 60°C, only E. faecalis will show growth.

Table 14-1 Examples of bacteria showing different temperatures for their growth Type of bacteria Psychrophilic Mesophilic Thermophilic

Example Psychrobacter immobilis, Methanosarcina spp. Escherichia coli. Thermus aquaticus, Bacillus stearothermophilus.

KEY FACTS 1 2 3 4

Temperature requirement for optimal growth and enzyme activity are often different. Different bacteria require different optimal temperature for their growth. Most of the medically important bacteria grow at 35°C -37°C. Campylobacter grows at 42°C.

VIVA 1 How does temperature affect bacterial growth? Ans: Bacterial growth is dependent on enzyme activity. Enzyme activity in turn is influenced by temperature. At high temperatures, they are denatured, while at low temperatures, they tend to get inactivated. Each species has an optimum range of temperature within which its reproductive rate and hence growth is optimum. 2 Give 2 examples each of psychrophilic, mesophilic and thermophilic bacteria. 3 Name some other enzymatic activities of the bacteria that are affected by temperature. Ans: Pigment production, production of flagella, and expression of stress proteins.

FURTHER READINGS 1 Bhattacharya S, Vijayalakshmi N, Parija SC. Uncultivable bacteria: Implications and recent trends towards identification. Indian J Med Microbiol. 2002: 20; 174-177. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 3 Isenberg HD. (Ed) Clinical Microbiology Procedures Handbook. American Society for Microbiology, Washington, DC, 1992. 4 PHLS Standard Operating Procedures. Inoculation of Culture Media. No B.SOP 54 Version: 1, 1998. 5 WHO. Guidelines on standard operating procedures for Microbiology. Blood safety and clinical technology. Chapter 6: Cultivation of bacteria on laboratory media. (World Health Organisation, Geneva) 1997.

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LESSON

pH Requirement for Growth of Bacteria

15 15

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to understand: 1 The pH requirement of bacteria.

1 Using sterile pipettes, inoculate 0.1 ml of each organism into nutrient broth tubes of pH, 3, 5, 7 and 9. 2 Incubate the tubes for 24-48 hr. 3 Check for growth of bacteria in these tubes.

INTRODUCTION Like temperature, pH also plays an important role in the growth and reproduction of bacteria. Each species of bacteria can grow within a particular pH range, and maximum growth occurs in an optimum pH range. This is a reflection of this natural environment eg. enteric bacteria such as Escherichia coli have a broad pH range, similar to that of the gut. Generally, bacteria grow best at a pH between 5.5-8 optimum pH being 6.5-7.5 Fungi thrive in an acidic environment of about pH 4-6. Most laboratory media have a neutral pH which suits nearly all organisms.

QUALITY CONTROL 1 Nutrient broth tube inoculated with Candida spp as quality control for growth at acidic pH. 2 Nutrient broth tube inoculated with E. coli as quality control for growth at pH 7.2 3 Nutrient broth tube inoculated with A. faecalis as quality control for growth at alkaline pH.

OBSERVATIONS PRINCIPLE Since pH changes can occur during growth of the bacteria due to the accumulation of metabolic byproducts, buffers are incorporated into the medium to prevent pH change. These may include natural buffers like proteins, peptone and amino acids which retard the shift because of their amphoteric nature. Salts of weak acids and weak bases may also be added.

REQUIREMENTS I Equipments Bacteriological incubator. II Reagents Nutrient broth tubes, 3 each of pH, 3, 5, 7 and 9. III Specimen A 24 hour broth culture of Escherichia coli, Alcaligenes faecalis and Candida spp.

1 Reproduction of bacteria is maximal at optimal pH. 2 Ability to tolerate and grow at a particular pH will be seen as turbidity in culture tubes.

RESULTS AND INTERPRETATION 1 All 3 microorganisms (E. coli, A. faecalis and Candida spp) show growth and turbidity at pH 7. 2 No microorganism shows any growth at pH 3. 3 E. coli and A. faecalis do not show any growth at pH 3 or pH 5. 4 Only A. faecalis shows growth and turbidity at pH 9. 5 Only Candida spp shows growth and turbidity at pH 5. The ability to grow at different pH varies among bacteria. Most pathogens show an optimum pH close to that of their preferred habitat.

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pH Requirement of Growth of Bacteria

KEY FACTS 1 pH is a critical factor affecting growth and metabolism of bacteria. 2 Most laboratory media has a pH of 7 and incorporate buffers for maintaining the same. 3 Extremes of pH can be used to selectively grow some microorganisms.

VIVA 1 Name some buffers used in preparing culture media. Ans: Citrate buffer, bicarbonate buffer and phosphate buffer. 2 Name bacteria that can be selectively grown in alkaline medium. Ans: Vibrio cholerae and Enterococcus faecalis. 3 Why does optimum growth of most pathogenic bacteria occur at pH 7.2? Ans: Most body fluids have a pH of 7.2. Pathogenic bacteria have adapted to their hosts, and hence are best capable of survival at a pH identical to that of the host.

FURTHER READINGS 1 Bhattacharya S, Vijayalakshmi N, Parija SC. Uncultivable bacteria: Implications and recent trends towards identification. Indian J Med Microbiol. 2002: 20; 174-177. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 3 Isenberg HD. (Ed) Clinical Microbiology Procedures Handbook. American Society for Microbiology, Washington, DC, 1992. 4 PHLS Standard Operating Procedures. Inoculation of Culture Media. No B.SOP 54 Version: 1, 1998. 5 WHO. Guidelines on standard operating procedures for Microbiology. Blood safety and clinical technology. Chapter 6: Cultivation of bacteria on laboratory media. (World Health Organisation, Geneva) 1997.

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LESSON

16

Oxygen Requirement for Growth of Bacteria

LEARNING OBJECTIVES After completing this practical you will be able to understand: 1 The oxygen requirement of bacteria.

INTRODUCTION Oxygen is one of the most important growth limiting factor for microorganism. It plays a vital role in many biological processes of the microorganisms. Oxygen requirement vary widely among bacteria and this is reflective of the different bio-oxidative enzyme systems present in bacterial cells.

pathway is followed in absence of oxygen substrate when substances like nitrate, sulphates, etc. can act as terminal electron acceptor. When inoculated into a tube of semisolid or liquid medium, aerobes grow only on the surface while obligate anaerobes grow only in the depths of the tube. Facultative anaerobes are evenly dispersed throughout the medium while microaerophiles grow slightly below the surface (subsurface growth). Some bacteria require the presence of 5 to 10% C02 for better growth of the bacteria by using candle jar (Fig. 16-1).

REQUIREMENTS PRINCIPLE On the basis of oxygen requirement, bacteria can be classified into 5 groups as follows: 1 Aerobes: These bacteria can grow only in the presence of free oxygen. Enzyme systems present in these bacteria require oxygen to be the final electron (hydrogen) acceptor especially for the oxidative breakdown of high energy molecule like glucose. 2 Microaerophiles: These bacteria require small amount of oxygen for their growth and survival. An excess of oxygen is inhibitory to their growth. 3 Obligate anaerobes: These bacteria cannot survive in the presence of free oxygen, hence other molecule act as the final electron acceptor. 4 Aerotolerant anaerobes: They do not use oxygen as a final electron acceptor but possess enzymes like superoxide dismutase and catalase, which can prevent the accumulation of toxic metabolites produced in the presence of oxygen. Therefore, oxygen is not lethal to them. 5 Facultative anaerobes: These bacteria can grow in the presence or absence of free oxygen. They generally follow an aerobic respiratory pathway, an anaerobic respiratory

I Equipments Bacteriological incubator and water bath. II Reagents and media Nutrient broth tubes and brain heart infusion (BHI) agar tubes. III Specimen A 24 hour nutrient broth culture of Escherichia coli, Pseudomonas aeruginosa, and 48 hour thioglycollate broth culture of Clostridium sporogenes.

FIGURE 16-1 Candle jar used for incubation and culture of bacteria..

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Oxygen Requirement for Growth of Bacteria

PROCEDURE 1 Warm the BHI agar tube in a water bath so that the agar melts. 2 Cool the medium to 45°C. 3 With a sterile inoculating loop, introduce a loopful of the culture into appropriately labeled tube. 4 Rotate the tubes between palms to evenly dispense the organisms. 5 Incubate the tubes for 24 hours at 37°C. 6 Innoculate thioglycollate broth with Cl. sporogenes and incubate at 37°C for 48 hours.

QUALITY CONTROL 1 BHI agar tube inoculated with P. aeruginosa, the bacteria that does not grow anaerobically 2 Thioglycollate broth culture inoculated with Cl. sporogenes, the bacteria that does not grow aerobically

OBSERVATIONS 1 Bacteria will grow at different depths in the tube depending on their oxygen requirement. Aerobes will grow on the surface of the medium, anaerobes in the depth of the medium,

and facultative anaerobes throughout the medium in the tube (Table 16-1). 2 Growth will be seen as turbidity in the appropriate section of the tube.

RESULTS AND INTERPRETATION Ps. aeruginosa will grow only on the surface of the medium. E. coli will grow throughout the tube and Cl. sporogenes only in the depths at the bottom of the tube. Ps. aeruginosa is an aerobe, Cl. sporogenes is an obligate anaerobe and E. coli is a facultative anaerobe.

Table 16-1 Examples of bacteria grouped depending on their requirement of oxygen Type

Example

Obligate aerobes Microaerophiles Obligate anaerobes Aerotolerant anaerobes Facultative anaerobes

Vibrio cholerae Clostridium perfringens Clostridium tetani Clostridium histolyticum Escherichia coli

KEY FACTS 1 Oxygen is both beneficial and toxic to living organisms. The benefits are because of their ability to act as a final electron acceptor in the respiratory pathway with the liberation of much energy. However it also results in the accumulation of toxic byproducts. 2 Bacteria may possess enzyme system to remove these toxic substances and thus help in their survival in the presence of oxygen. 3 Anaerobic bacteria can be cultured by the exclusion of air using anaerobic jars, prereduced media, etc.

VIVA 1 Give example of obligate aerobes, microaerophiles, obligate anaerobes, aerotolerant anaerobes and facultative anaerobes. 2 What are the media commonly used to culture anaerobes? Ans: Robertson’s cooked meat (RCM) medium, thioglycollate broth, blood agar incorporating antibiotics like neomycin or kanamycin.

FURTHER READINGS 1 Bhattacharya S, Vijayalakshmi N, Parija SC. Uncultivable bacteria: Implications and recent trends towards identification. Indian J Med Microbiol. 2002: 20; 174-177. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 3 Isenberg HD. (Ed) Clinical Microbiology Procedures Handbook. American Society for Microbiology, Washington, DC, 1992. 4 PHLS Standard Operating Procedures. Inoculation of Culture Media. No B.SOP 54 Version: 1, 1998. 5 WHO. Guidelines on standard operating procedures for Microbiology. Blood safety and clinical technology. Chapter 6: Cultivation of bacteria on laboratory media. (World Health Organisation, Geneva) 1997.

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LESSON

Culture of Anaerobic Bacteria

17 17

LEARNING OBJECTIVES After completing this practical you will be able to understand: 1 The methods of culture of anaerobic bacteria.

INTRODUCTION Obligate anaerobes can grow in media only in the absence of oxygen, ie. from which air is excluded. These organisms die rapidly on exposure to air, hence for maintaining anaerobiosis various methods have been devised for anaerobic culture.

PRINCIPLE There are different ways of creating anaerobic conditions suitable for the growth of obligate anaerobes. Deep nutrient agar tubes are the simplest method. The tubes are inoculated while still molten, cooled rapidly and incubated. Anaerobes grow in the depths of the medium, and the number of colonies becomes fewer towards the surface. Strict anaerobes will not grow within a centimeter of the surface. Alternatively, reducing agents like 0.5-1% glucose, 0.1% ascorbic acid, 0.1% cysteine, 0.1% thioglycollate can be added. Cooked meat particles also act as a good reducing agent Example. Robertson Cooked meat medium (Fig. 17-1). For culture of anaerobes, oxygen must be excluded either by combustion or by replacing it with an inert gas. In many laboratories, combustion involves the combining of oxygen with hydrogen to form water in the presence of a catalyst like palladium or palladinized asbestos. Anaerobic jars are a constant feature of anaerobic culture. They include the McIntosh and Fildes jar (Fig. 17-2) , which has inlets to admit hydrogen and carbon dioxide, a vacuum pump for evacuating oxygen, and a catalyst fitted into the lid. A simpler but more expensive technique is the Gaspak system. This utilizes a transparent polycarbonate jar with a lid

bearing a screened catalyst chamber. The catalyst, consisting of pellets of sodium borohydride, cobalt chloride, citric acid and sodium bicarbonate is contained in sachets. Water is added to the sachet and it is immediately placed in the jar, which is then sealed tightly. The resulting reaction liberates hydrogen and carbon dioxide. An indicator is also added to demonstrate anaerobiosis. List of anaerobic bacilli and cocci are summarized in the box 17-1.

REQUIREMENTS I Equipments Anaerobic culture systems: McIntosh and Fildes jar and Gaspak system. II Reagents and media Blood agar plates and thioglycollate broth culture. III Specimen A 48 hour thioglycollate broth culture of anaerobic bacteria (Clostridium sporogenes, Bacteroides spp), and 24 hour cultures of facultative anaerobes (Escherichia coli), and obligate aerobes (Pseudomonas aeruginosa).

PROCEDURE 1 Divide each plate into 4 quadrants. 2 Inoculate a loopful of each organism into a quadrant. 3 Stack the plates into the anaerobic jars, introduce the catalyst and quickly seal the lid. Note: Anaerobic condition should be checked by alkaline methylene blue indicator. 4 Incubate the plates at 37ºC for 48 hours. 5 Incubate one plate aerobically. 6 Remove the plates from the jars and examine for growth.

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Culture of Anaerobic Bacteria

FIGURE 17-1 Robertson’s cooked meat medium.

QUALITY CONTROL 1 Blood agar inoculated with P. aeruginosa, the bacteria that does not grow anaerobically 2 Thioglycollate broth culture inoculated with Cl. sporogenes, the bacteria that does not grow aerobically

OBSERVATIONS If anaerobiosis is complete, obligate anaerobes like Cl. sporogenes will grow, while obligate aerobes like P. aeruginosa will not grow.

RESULTS AND INTERPRETATION P. aeruginosa will not show growth on the plates incubated anaerobically, while Cl. sporogenes and Bacteroides spp will grow on the blood gar plates. P. aeruginosa is a strict aerobe that cannot grow in the absence of oxygen. P. aeruginosa will show growth on the aerobically incubated plate while Cl. sporogenes or Bacteroides spp will not grow.

FIGURE 17-2 Macintosh Filde’s jar.

Cl. sporogenes and Bacteroides are strict anaerobes that cannot grow if oxygen is present. E. coli will grow on all the plates, either incubated aerobically or anaerobically. E. coli is a facultative anaerobe and can grow either in the presence or absence of oxygen.

BOX 17-1 LIST OF ANAEROBIC BACILLI AND COCCI Anaerobic bacilli Gram positive bacilli Bifidobacterium Propionobacterium Eubacterium Lactobacillus Actonomyces Anaerobic cocci Gram positive cocci Peptosteptococcus Coprococcus Ruminococcus

Gram negative Bacilli Bacteroides Fusobacterium

Gram Negative cocci Veillonella Acidaminococcus Megasphaera

VIVA 1 List some methods of anaerobic culture not utilizing anaerobic jars. 2. What is the Hungate procedure of anaerobiosis? Ans. The Hungate procedure is a method for the anaerobic culture of bacteria. In the presence of an oxygen free gas, usually nitrogen, a thin layer of agar is coated on the inside of the culture tube. The organisms are then inoculated, also in the presence of the same gas. The tubes are then sealed. This ensures anaerobic conditions inside the tube.

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KEY FACTS 1 If anaerobiosis is inadequate, obligate anaerobes may fail to grow. 2 The addition of indicators to an anaerobic system is a useful way of ascertaining absence or presence of oxygen.

FURTHER READINGS 1 Bhattacharya S, Vijayalakshmi N, Parija SC. Uncultivable bacteria: Implications and recent trends towards identification. Indian J Med Microbiol. 2002: 20; 174-177. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 3 Isenberg HD. (Ed) Clinical Microbiology Procedures Handbook. American Society for Microbiology, Washington, DC, 1992. 4 PHLS Standard Operating Procedures. Inoculation of Culture Media. No B.SOP 54 Version: 1, 1998. 5 WHO. Guidelines on standard operating procedures for Microbiology. Blood safety and clinical technology. Chapter 6: Cultivation of bacteria on laboratory media. (World Health Organisation, Geneva) 1997.

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LESSON

18

Sterilization of Commonly Used Culture Media

LEARNING OBJECTIVES After completing this practical you will be able to become familiar with: 1 The commonly used techniques for the sterilization of culture media.

Chamber land),b) asbestos filters (e.g., Seitz filter (Fig. 18-1), c) sintered glass filter (Fig. 18-2), and d) cellulose membrane filters. Of these cellulose membrane filter is most extensively used now a days. Different methods of sterilization of various substances are summarized in the table 18-1.

REQUIREMENTS INTRODUCTION Sterilization is the process by which an article is freed of all living organisms, including spores. This is vital for isolation and maintenance of microbes. The common methods used for sterilization of media is heat and filtration.

I Equipments Autoclave (Fig. 18-3), hot air oven (Fig. 18-4) and Seitz filter. II Reagents Sterile nutrient broth, Browne’s tube no. 3, and filter paper strips impregnated with spore of Clostridium tetani and filter paper strips impregnated with spores of Bacillus stearothermophilus.

PRINCIPLE Sterilization by heat This can be performed by two methods as mentioned below: Dry heat: Sterilisation by dry heat kills the bacteria by oxidizing essential cell components of the cell. The hot air oven is commonly used in a microbiology laboratory for sterilization of laboratory glassware, oils, powders etc. In a hot air oven, temperatures maintained at 160°C, for an hour is effective for sterilization. Moist heat: Moist heat kills the microorganisms by coagulating their proteins and denaturing their enzymes. Moist heat has greater penetrating power than the dry heat and so relatively lower temperature is required for sterilization by this method. Pasteurisation is an example of sterilization by moist heat (Box 18-1).

Sterilization by filtration Heat labile fluids such as serum, antibiotics, etc. are sterilized by passing them through special filters. There are different types of filters. These include a) earthenware candles (e.g., Berkefield,

III Specimen Spore suspensions of Bacillus spp and suspension of Escherichia coli

PROCEDURE 1 Soak strips of filter paper in the spore suspensions of Bacillus spp and dry them in a Petri dish in the incubator. 2 Take 7 test tubes and place a dried strip in each of them. 3 Place the first three tubes in a hot air oven at 160°C for 20, 30 and 60 minutes respectively. 4 Autoclave the 4th, 5th and 6th tubes for 10, 20, and 30 minutes respectively. 5 After cooling, aseptically add 5 ml of nutrient broth to each tube and incubate for 24 hours at 37°C. 6 The 7th tube acts as control. 7 Inoculate a drop of the E.coli suspension onto a nutrient agar plate. 8 Filter the suspension through a Seitz filter. 9 Aseptically inoculate the filtrate onto a nutrient agar plate and incubate both plates at 37°C for 24 hours.

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FIGURE 18-1 Seitz filter.

QUALITY CONTROL 1 Autoclave: Filter paper strips impregnated with spores of B.stearothermophilus are autoclaved along with the normal load and then transferred to nutrient broth and incubated. If the correct sterilization conditions have been achieved no growth should occur. 2 Hot air oven: This can be tested by using a) Browne’s tube no. 3: Color change from red to green is a satisfactory result; b) Filter paper strips impregnated with spores of non toxigenic strain of C. tetani heated along with the load and incubated in appropriate media. No growth should occur. 3 Filters: Efficient filters should be able to retain Serratia marcescens.

FIGURE 18-3 Laboratory autoclave.

RESULTS AND INTERPRETATION 1 There will be growth in tubes, kept in the hot air oven for 20 minutes and 30 minutes but not for 60 minutes. 2 Only the tube autoclaved for 10 minutes will show growth. The other two tubes will remain free of growth. 3 There will be no growth in the culture filtrates. Both dry and moist heats are effective sterilization methods. However moist heat is more effective requiring less time and temperature. Filtration is also a suitable method of sterilization.

OBSERVATIONS If the spores have been killed by the heat process there will be no growth as demonstrated by a lack of turbidity. Similarly, if the bacteria have been held back by the filter there should be no growth in the sample of filtrate.

FIGURE 18-4 Hot air oven. FIGURE 18-2 Sintered glass filter.

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Sterilization of Commonly Used Culture Media

BOX 18-1 PASTEURISATION Pasteurization is a method of sterilisation by moist heat. It is sufficient to kill heat labile bacteria, but not spores. Two methods are available: holder method (60°C for 30 minutes) and flash method (72° C for 15 seconds). Some pathogens such as Coxiella burnetii is not destroyed, so this is not a very efficient method of sterilization.

Table18-1 Different methods of sterilization of various substances Methods of sterilization

Substances sterilized

Dry heat Flaming Hot air oven Incineration

Inoculating wires, loops. Glassware, oils, powders, paraffin . Biohazardous material- used gloves, needles, etc.

Moist heat Pasteurization Inspissation Boiling Tyndallization Autoclaving Filtration

Serum, milk. Löwenstein Jensen media, Loeffler’s serum slope. Needles, glass syringes. Sugar solution. Heat stable media such as nutrient agar. Heat labile media, serum, antibiotics, etc.

KEY FACTS 1 Various factors influence sterilization by heat. These include time, temperature, number of microorganisms and spores and their type, nature of material, etc. 2 It is important not to overfill the media container. 3 The load should be properly plugged or wrapped to ensure sterility. 4 The load should be cooled before being removed from the autoclave/hot air oven. 5 Filters should be assembled and autoclaved prior to use.

VIVA 1 Why should media not be sterilized in large quantities? Ans: Large quantities of media are difficult to sterilize by heat. It will not be possible to ensure that all parts of the media have attained sterilization temperature, and thus sterilization cannot be guaranteed. If methods like filtration are used, large volumes cannot be handled by the filtration apparatus. 2 What are the different methods of sterilizing by dry heat? Ans: These are sterilization by red heat, flaming, hot air oven and incineration. 3 How would you sterilize a) sera, b) Löwenstein Jensen medium, c) nutrient agar and d) paraffin?

FURTHER READINGS 1 Bhattacharya S, Vijayalakshmi N, Parija SC. Uncultivable bacteria: Implications and recent trends towards identification. Indian J Med Microbiol. 2002: 20; 174-177. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 3 Isenberg HD. (Ed) Clinical Microbiology Procedures Handbook. American Society for Microbiology, Washington, DC, 1992. 4 PHLS Standard Operating Procedures. Inoculation of Culture Media. No B.SOP 54 Version: 1, 1998. 5 WHO. Guidelines on standard operating procedures for Microbiology. Blood safety and clinical technology. Chapter 6: Cultivation of bacteria on laboratory media. (World Health Organisation, Geneva) 1997.

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LESSON

Antiseptics and Disinfectants

19 17

LEARNING OBJECTIVES After completing this practical you will be able to become familiar with: 1 The commonly used antiseptics and disinfectants and methods of testing them.

II Reagents and media Sterile nutrient broth, nutrient agar plates, used as well as fresh disinfectant (e.g. lysol). III Specimen A 24 hour broth culture of Escherichia coli.

PROCEDURE INTRODUCTION Disinfection is defined as the destruction of microorganisms not including bacterial spores. The process reduces microorganism to a level acceptable for a defined purpose. A chemical agent that disinfects a substance is called a disinfectant. A disinfectant that can be safely applied to living tissue is called an antiseptic. Table 19-1 summarises the list of commonly used disinfectants and antiseptics.

1 With a sterile pipette, transfer 1 ml of the used disinfectant into 9 ml sterile nutrient broth. 2 Also mix 1 ml of fresh disinfectant with 9 ml of E coli culture. 3 Inoculate this mixture onto 10 different areas of two well dried nutrient agar plates each. 4 Incubate one plate for 3 days at 37°C and the other for 7 days at room temperature.

QUALITY CONTROL PRINCIPLE Disinfectants are used for medical devices where sterility is not required. They have markedly different activity against different microorganisms and are generally most effective against Gram positive bacteria. The concentration at which they are used should be accurate for optimal activity. The article to be disinfected should be cleaned thoroughly (to reduce organic matter which may inactivate the agent) or immersed in the disinfection for the required amount of time. Commonly used disinfectants and their mechanism of actions are summarized in the box 19-1.

REQUIREMENTS I Equipments Sterile pipette.

1 Fresh disinfectant should be a) sterile, and b) should kill a test inoculum of E coli within 10 minutes.

OBSERVATION If the disinfectant has lost activity it will fail to kill microorganism on objects immersed in it. This will show a growth on the nutrient agar plate.

RESULTS AND INTERPRETATION If growth occurs on 5 or more spots on either plate, the used disinfectant is said to have failed the test. With time and use, the activity of a disinfectant decreases. The fresh disinfectant will be able to bring about disinfection more effectively than the used one.

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Table 19-1 List of commonly used disinfectants and antiseptics Disinfectants Lysol Zephiran Glutaraldehyde Sodium hypochlorite

Antiseptics 70% ethyl alcohol Chlorhexidine Iodine Dettol

BOX 19-1 COMMONLY USED DISINFECTANTS AND THEIR MECHANISM OF ACTIONS 1 Phenols (e.g. lysol, chloroxylenol): They act by a number of mechanisms such as disruption of cells, precipitation of protein inactivation of enzyme, etc. Phenol is the standard disinfectant against which other disinfectants are compared. 2 Alcohols (e.g. ethyl alcohol, usually at a concentration of 70%): They can also be used as antiseptics and are active against viruses as well. They act by denaturing proteins damaging lipid complexes and dehydration. 3 Halogens (eg. iodine, chlorine and sodium hypochlorite): Iodine is commonly used as an antiseptic. It acts by oxidation. Chlorine is a widely used disinfectant. The compounds act by the liberation of nascent oxygen and are effective against most bacteria and viruses. 4 Heavy metals and their compounds (e.g. copper sulphate): They act by inactivating cellular protein. 5 Synthetic detergents (e.g. sodium lauryl sulfate): They help in mechanical removal of microorganism. 6 Quaternary ammonium compound (e.g. zephiran): They act on the cell membrane of bacteria.

KEY FACTS 1 2 3 4

Disinfectant does not guarantee sterility. They merely ensure that an object is relatively free of microbial contamination. Antiseptics can be used for decontamination of living tissue. Disinfectants are used for decontamination of non-living tissue. It is essential to test disinfectant from time to time to detect loss of activity.

VIVA 1 What are the commonly used disinfectants and antiseptics? 2 What is the difference between sterilization and disinfection? Ans: Sterilization frees an article of all infectious materials including spores, whereas disinfection merely frees an article of infectious, vegetative cells. It doesn’t guarantee the removal or inactivation of spores. In general, the temperatures used in sterilization are higher than those used for disinfection, and the concentration of chemical agents is higher when they are used for sterilization than disinfection. 3 What are different methods used for testing disinfectant? Ans: These are : a) Rideal Walker test, b) Chick Martin test and c) In-use test. FURTHER READINGS 1 Bhattacharya S, Vijayalakshmi N, Parija SC. Uncultivable bacteria: Implications and recent trends towards identification. Indian J Med Microbiol. 2002: 20; 174-177. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 3 Isenberg HD. (Ed) Clinical Microbiology Procedures Handbook. American Society for Microbiology, Washington, DC, 1992. 4 PHLS Standard Operating Procedures. Inoculation of Culture Media. No B.SOP 54 Version: 1, 1998. 5 WHO. Guidelines on standard operating procedures for Microbiology. Blood safety and clinical technology. Chapter 6: Cultivation of bacteria on laboratory media. (World Health Organisation, Geneva) 1997.

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UNIT

IV ENZYMATIC AND BIOCHEMICAL ACTIVITIES OF BACTERIA Lesson 20 Catalase Test Lesson 21 Oxidase Test Lesson 22 Coagulase Test Lesson 23 Urease Test Lesson 24 Indole Test Lesson 25

Methyl Red Test

Lesson 26 Voges-Proskauer Test Lesson 27 Citrate Utilization Test Lesson 28 Triple Sugar Iron (TSI) Agar Test Lesson 29 Hydrogen Sulphide Test Lesson 30

Nitrate Reduction Test

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LESSON

20

Catalase Test

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to:

I Reagents and glass wares 3% hydrogen peroxide, glass slides, test tubes, glass rod / platinum loop / plastic loop and other standard lab wares.

1 Demonstrate the presence of catalase, an intracellular enzyme in the bacteria. 2 Distinguish the bacteria based on the catalase activity.

INTRODUCTION Catalase is an enzyme produced by many bacteria. The enzyme splits hydrogen peroxide into water and oxygen. Hydrogen peroxide is a by product of aerobic respiration and is lethal if it accumulates in the bacterial cell. Catalase degrades the hydrogen peroxide in the bacterial cell before it can do any damage to the bacterial cell.

II Specimen Pure growth of bacteria from solid media preferably from nonblood agar plates (Examples: nutrient agar, Muller-Hinton agar) is tested.

PROCEDURE Test can be done by 2 methods as follows: 1 Slide method 2 Tube method

PRINCIPLE Slide method Chemically, catalase is a haemoprotein, similar in structure to haemoglobin, except that the four iron atoms in the molecule are in the oxidized (Fe3+) rather than the reduced (Fe2+) state. The enzyme converts hydrogen peroxide into water and oxygen.

1 Transfer pure growth of the organism from the agar to a clean slide with a loop or glass rod. 2 Immediately add a drop of 3% hydrogen peroxide to the growth. 3 Observe for bubble formation.

2H202 —— 2H2 0+02 (gas bubbles)

Tube method Production of the enzyme catalase can be demonstrated by adding hydrogen peroxide to colonies of the bacteria. If catalase is present it is indicated by the presence of free gas bubbles. If catalase is absent, no bubbles will be seen. The catalase test is most commonly used to differentiate members of the family Micrococcaceae from members of the family Streptococcaceae (Box 20-1). Catalase test is also carried out for Mycobacteria to differentiate tubercle bacilli from atypical mycobacteria (Box 20-2). List of catalase positive and negative bacteria are summarized in the table 20-1.

1 Take 1 ml of 3% hydrogen peroxide in 12 x 100 mm test tube. 2 Introduce small quantity of bacterial growth into the fluid with the help of a glass rod or plastic loop and touch the side of the tube. 3 Observe the release of bubbles. Note: Hydrogen peroxide must be stored in amber coloured bottles. If stored in colourless penicillin vials, the hydrogen peroxide on exposure to light will be broken down into oxygen and water. Use of this solution will give false negative results.

Textbook of Practical Microbiology

QUALITY CONTROL Positive control: Staphylococcus aureus (catalase positive bacteria). Negative control: Streptococcus species (catalase negative bacteria).

OBSERVATIONS

Table 20-1 Catalase positive and negative bacteria Catalase positive bacteria

Catalase negative bacteria

Staphylococci Micrococci Corynebacterium diphtheriae Enterobacteriaceae

Streptococcus pyogenes Gardnerella vaginalis Fusobacterium species Eikenella corrodens Kingella kinge Shigella dysenteriae type 1 Fatumella ptysees

Slide method Gas bubbles are formed immediately when 3% H2 02 is added to the colony. Tube method Gas bubbles are released when colonies are introduced into the hydrogen peroxide in the test tube.

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BOX 20-1 USES OF CATALASE TEST The catalase test is useful to differentiate:

RESULTS AND INTERPRETATION 1 The rapid and sustained appearance of bubbles or effervescence constitutes a positive test. It means bacteria possesses the enzyme catalase, hence is catalase positive. 2 Some bacteria possess enzymes other than catalase that can decompose hydrogen peroxide. Hence, forming a few tiny bubbles after 20-30 seconds is not considered a positive test.

1 Staphylococci (catalase +ve) from streptococci and enterococci (catalase - ve). 2 Listeria monocytogenes and Corynebacteria (catalase +ve) from other Gram positive, non-spore forming bacilli (catalase - ve). 3 Members of family Enterobacteriacae. 4 Aerobic bacteria (catalase +ve) from anaerobic bacteria (catalase - ve).

BOX 20-2 CATALASE TEST FOR MYCOBACTERIA It helps to differentiate tubercle bacilli from atypical mycobacteria. Most atypical mycobacteria are strongly catalase positive, while tubercle bacilli are weakly positive. Most strains of mycobacteria, except certain strains of Mycobacterium tuberculosis complex (some isoniazid resistant strains) and M. gastri, produce the intracellular enzyme catalase. The test is performed by mixing equal volumes of 30% hydrogen peroxide and 0.2% catechol in distilled water and then adding it to 5ml of a mycobacterial test culture. It is allowed to stand for a few minutes. Effervescence indicates catalase production. Catalase production is assessed by a) Relative activity of the enzyme catalase determined by the height of the column of oxygen bubbles formed by the action of untreated enzyme produced by the organism (Semi quantitative catalase test). On the basis of the semi quantitative catalase test, mycobacteria are classified in to 2 groups: i. those producing 45mm column height of bubbles. b) By the ability of the enzyme catalase to remain active after heating, a measure of the heat stability of the enzyme (Heat stable catalase test). When heated to 68°C for 20 minutes, the catalase of M. tuberculosis. M. bovis, M. gastri and M. haemophilum becomes inactivated.

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KEY FACTS 1 2 3 4 5 6

Catalase enzyme splits hydrogen peroxide into oxygen and water. In aerobic organisms, 02 serves as H2 acceptor and hydrogen peroxide (H2 02) is formed in the cell. High concentration of H2 02 is toxic to the cell. Therefore, aerobic bacteria possess catalase to split toxic hydrogen peroxide. Most of the aerobes are catalase positive. Most of the anaerobes do not possess the enzyme, catalase. Care must be taken while performing catalase test in growth from blood agar plate. Blood (RBC) contains catalase, which gives false positive reactions.

VIVA 1 2 3 4 5

What is catalase test? What are the positive and negative controls used in the catalase test? What is the importance of catalase test? Can you take colonies from blood agar plate for testing catalase reaction? Give explanations. Name some catalase positive and negative organisms.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

Oxidase Test

21 17

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the presence of oxidase, an intracellular enzyme in the oxidase-positive bacteria. 2 Distinguish the bacteria based on the cytochrome oxidase activity.

dihydrochloride (1%), and dimethyl – p – phenylene diamine dihydrochloride (1%). Wood stick/platinum loop/glass rod, and filter paper. II Specimen Young culture of bacteria to be tested, preferably less than 24 hours old, growing on an agar plate or agar slant.

INTRODUCTION

PROCEDURE

The enzyme oxidase plays a vital role in the operation of the electron transport system during aerobic respiration. Cytochrome oxidase catalyzes the oxidation of a reduced cytochrome by molecular oxygen, resulting in the formation of water or hydrogen peroxide.Aerobic bacteria, as well as some facultative anaerobes and microaerophiles exhibit oxidase activity.

The test is performed by following two methods:

PRINCIPLE The cytochromes are iron containing haemoproteins that act as the last link in the chains of aerobic respiration by transferring electrons (hydrogen) to oxygen, with the formation of water. The cytochrome oxidase test uses certain reagent dyes such as p-phenylene diamine dihydrochloride which acts as a substitute for oxygen as artificial electron acceptors. This enzyme oxidises the reagent N-N tetramethyl para-phenylene diamine hydrochloride (a colour less reagent in reduced form) to indophenol blue, a purplish blue coloured product. List of oxidase positive and negative bacteria is presented in the table 21-1.

REQUIREMENTS I Reagents and glass wares Fresh reagent: Tetramethyl – p-phenylene diamine

1 Direct plate technique, and 2 Indirect paper strip procedure.

Direct plate technique 1 Take a nutrient agar plate with colonies of bacteria to be tested. 2 Add 2 to 3 drops of reagent (tetramethyl p-phenylene diamine hydrochloride or dimethyl-p- phenylene diamine dihydrochloride) directly to the bacterial colonies growing on medium in the plate. 3 Note the change of colour of the colonies.

Indirect filter paper strip procedure 1 Take a filter paper strip. 2 Moisten the filter paper strip with freshly prepared 1% oxidase reagent. Note: Oxidase reagent is freshly prepared in distilled water every day. 3 Pick up the colonies to be tested with the help of a glass rod or plastic loop or platinum wire. 4 Smear the colonies into the reagent zone of the filter paper. 5 Note the change in colour if any within 10 seconds.

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Oxidase Test

QUALITY CONTROL

Table 21-1 List of oxidase positive and negative bacteria

Positive control: Pseudomonas aeruginosa (oxidase positive bacteria). Negative control: Escherichia coli (oxidase negative bacteria).

Oxidase positive bacteria Gram negative rods 1. Pseudomonas spp(except Ps. cepacia). 2. Vibrio spp. 3. Aeromonas spp. 4. Camphylobacter spp. 5. Plesiomonas spp. 6. Flavobacterium spp. 7. Alcaligenes spp. 8. Haemophilus spp. 9. Moraxella spp. 10. Chromobacterium spp. 11. Bordetella spp(except B.parapertusis) 12. Brucella spp (except B.canis) 13. Eikinella spp. 14. Cardiobacterium spp. 15. Achromobacter spp. 16. Pasteurella multocida

OBSERVATIONS Direct plate technique In a positive test, bacterial colonies on the plate develop a deep blue colour at the site of inoculation within 10 seconds. In a negative test the colour of the colonies remain unchanged. Indirect filter paper strip procedure In a positive test, a deep blue colour develops at the site of smear in the filter paper, within 10 seconds (Fig. 21-1). In a negative test the colour of the smear in the zone of the filter paper remain unchanged.

RESULTS AND INTERPRETATION Bacterial colonies having cytochrome oxidase activity develop a deep blue colour at the inoculation site within 10 seconds. In filter paper test, deep blue colour develops at the site of smear within 10 seconds. It means bacteria possesses the enzyme oxidase, hence is oxidase positive.

Gram negative cocci 1. Neisseria spp. 2. Branhamella spp. Oxidative negative bacteria 1. All genera in family Enterobacteriaceae 2.Acinetobacter calcoaceticus 3. Bordetella parapertusis 4.Brucella canis 5.Francisella tularensis 6.Gardnerella vaginalis

FIGURE 21-1 Slide oxidase test

VIVA 1 What is the principle of oxidase test? 2 How is oxidase test done? 3 What are the precautions to be observed while doing oxidase test? Ans: a.Fresh reagent must be used. b. Iron wire loops should not be used. c. Nichrome wire loops should not be used. d. Colonies should not be picked up from selective media. e. Results must be observed within 10 seconds. 4 Name oxidase positive bacteria. 5 Name oxidase negative bacteria. 6 What is chemical name of the oxidase reagent?

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KEY FACTS 1 2 3 4

The dye, p-phenylene diamine dihydrochloride is the substitute for oxygen as artificial election acceptors. In reduced state the dye is colourless and in oxidized state deep purple in colour. Colour change must be noted within 10 seconds. Tetramethyl derivative of p-phenylene diamine is recommended because the reagent is i more stable in storage, ii more sensitive to the detection of cytochrome oxidase, and iii. less toxic than dimethyl derivative. 5 Nichrome or stainless steel inoculating loops or wires should not be used for performing the test because surface oxidation products formed during the process of sterilization by flaming may result in false positive reaction. 6 Always freshly prepared reagent should be used. 7 Colonies for testing should not be taken from blood agar.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

22

Coagulase Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the presence of coagulase, an intracellular enzyme in the bacteria, especially Staphylococcus aureus. 2 Distinguish the bacteria based on the coagulase activity.

by tube coagulase test. In this method, a suspension of coagulase producing staphylococci is prepared in plasma in a test tube, and incubated at 37°C for 3-6 hours. In a positive test, the enzyme coagulase secreted by S. aureus is liberated to the medium, which reacts with fibrinogen to produce a visible fibrin clot. List of coagulase positive bacteria is presented in the table 22-1.

INTRODUCTION REQUIREMENTS The enzyme, coagulase, produced by a few Staphylococcus species, is a key feature of pathogenic Staphylococcus. The enzyme causes coagulation of blood, allowing the organism to “wall” its infection off from the host’s protective mechanisms rather effectively. Coagulase is a protein having a prothrombin-like activity capable of converting fibrinogen into fibrin, which results in the formation of visible clot. In the laboratory, the coagulase test is used to identify S. aureus and differentiate it from the other species of coagulase-negative Staphylococcus.

I Reagents and lab wares Rabbit plasma with EDTA anticoagulant, saline, glass slides, test tubes, glass rod/platinum loop/plastic loop and other standard lab wares. II Specimen Pure growth of S. aureus from solid media preferably from nonblood agar plates (Examples: nutrient agar, Muller-Hinton agar).

PROCEDURE PRINCIPLE Slide test S. aureus produces the enzyme coagulase in 2 forms: a. bound coagulase and b. free coagulase.

Bound coagulase Bound coagulase is also known as clumping factor. It is bound to the bacterial cell wall and is not present in culture filtrates. Presence of this enzyme is tested by slide coagulase test. Fibrin strands are formed between the bacterial cells when suspended in plasma (fibrinogen), causing them to clump into visible aggregates.

1 Take a clean glass slide. 2 Mark it into two halves by a glass marking pencil. 3 Add two drops of sterile saline on two halves of the glass slides. 4 Pick up the colonies of S. aureus to be tested from agar culture and gently emulsify with drops of saline. 5 Add a drop of undiluted plasma to the bacterial suspension and mix with a wooden applicator sticks. 6 Place another drop of saline in other half of the slide as a control. 7 Rock the slide, back and froth, and observing for the prompt clumping of the bacterial suspension within 10-15 seconds.

Free coagulase Tube test Free coagulase is a thrombin-like substance present in S. aureus culture filtrates (Box 22 -1). Presence of free coagulase is tested

1 Take 0.5 ml of rabbit plasma (diluted 1 in 5 with saline) in a test tube.

Textbook of Practical Microbiology

2 Add approximately 5 drops (250 µl) of overnight broth culture or small amount of the colony growth of S. aureus to the diluted plasma in the test tube. 3 Incubate the tube at 37°C for 4 hours. 4 Observe for clot formation by gently tilting the tube. 5 If no clot is observed at that time, reincubate the tube at room temperature and read again after 18 hours.

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POSITIVE

QUALITY CONTROL Positive control: S. aureus (Coagulase positive bacteria). Negative control: S. epidermidis (Coagulase negative bacteria). Coagulability of plasma may be tested by adding one drop of 5% calcium chloride to 0.5 ml of the reconstituted plasma. A clot should form within 10 to 15 seconds.

OBSERVATION In a positive slide test, prompt clumping of the organism shows the presence of the bound coagulase. In a positive tube test, the plasma in the tube clots and does not flow when the tube is inverted (Fig. 22-1). Note: On continued incubation, the clot may be lysed by fibrinolysin secreted by some strains.

RESULTS AND INTERPRETATION In slide test, Positive reaction will be detected within 10–15 seconds of mixing the plasma with the suspension by the formation of a white precipitate and agglutination of the

NEGATIVE FIGURE 22-1 Tube coagulase test.

organisms. The test is considered negative if no agglutination is observed after 2 minutes. All strains that are coagulase positive can be reported as S. aureus. All strains producing negative slide tests must be tested with the tube coagulase test. The tube coagulase test is considered positive if any degree of clotting is noted.

Table 22-1 List of coagulase positive bacteria 1 2 4 5 6 7 8

Staphylococcus aureus. Staphylococcus schleiferi Staphylococcus felis Staphylococcus lutrae Staphylococcus intermedius Staphylococcus hyicus Peptostreptococcus hydrogenalis

BOX 22-1 FREE COAGULASE Free coagulase is an extracellular enzyme produced by Staphylococcus aureus. It activates a coagulase reacting factor (CRF) normally present in the plasma to clot by the conversion of fibrinogen to fibrin. Coagulase does not clot plasma of guinea pigs because they lack CRF. There are 8 antigenic types of coagulase (designated as A to H).Most human strains of S. aureus produce coagulase A. All coagulase producing staphylococci are S. aureus and as a result coagulase production is considered the best laboratory evidence for the potential pathogenicity of Staphylococcus. Coagulase may act to coat the bacterial cells with fibrin rendering them resistant to opsonization and phagocytosis.

KEY FACTS 1 2 3 4 5 6

Two types of coagulase are produced by S. aureus. These are bound coagulase, and tube coagulase. Bound coagulase is tested by slide coagulase test. Free coagulase is tested by tube coagulase test. In the tube coagulase test, on continued incubation, the clot already formed may be lysed by fibrinolysin secreted by some strains. Rabbit plasma with EDTA is used in both tests. Pooled human plasma can be used after checking with a standard strain of S. aureus.

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VIVA 1 2 3 4 5 6 7

What is the principle of slide coagulase test? What is the principle of tube coagulase test? What is the positive control in the test? What is the negative control in the test? How will you interpret the slide coagulase test? How will you interpret the tube coagulase test? Give examples of coagulase positive bacteria.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

23

Urease Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the presence of urease, an intracellular enzyme in the bacteria. 2 Distinguish the bacteria based on the urease activity.

II Reagents and glass wares Inoculating wire, Christensen’s urea agar, and 12 × 100 mm test tubes. III Specimen Pure growth of Proteus mirabilis from solid media preferably from non-blood agar plates (Examples: nutrient agar, MullerHinton agar) is tested.

INTRODUCTION PROCEDURE Certain bacteria and fungi possess the enzyme urease that hydrolyzes urea releasing ammonia into the medium. This produces a change in the pH of the medium that can be detected by the color change in the indicator dye. This test can be used to differentiate different groups of bacteria and fungi.

1 Pick up the colonies of P. mirabilis from the culture on nutrient agar. 2 Inoculate Christensen’s urea agar slope with these bacterial colonies. 3 Incubate the tube at 37°C for 18 hours. 4 Observe any change of colour in the inoculated medium.

PRINCIPLE Urea is a diamide of carbonic acid. Urease, the enzyme produced by the bacteria and fungi, hydrolyses urea and releases ammonia and carbon dioxide. Ammonia reacts in solution to form ammonium carbonate, which is alkaline leading to an increase in pH of the medium. Phenol red that is incorporated in the medium changes its color from yellow to red in alkaline pH, thus indicating the presence of urease activity. The composition and preparation of Christensen’s urea agar is described in the box 23-1. List of urease producing microorganisms is summarized in the Table 23-1.

REQUIREMENTS I Equipments Incubator.

BOX 23-1 CHRISTENSEN’S UREA AGAR Composition Peptone – 0.1 gm. Glucose – 0.1 gm. NaCl – 0.5 gm. Monopotassium phosphate – 0.2 gm. Phenol red (1.2%) – 1.0 ml. Agar – 2 gm. Distilled water – 100 ml. pH – 6.8. Preparation Prepare the base, sterilize by autoclaving at 121°C for 15 min. Cool to 50°C in a waterbath and then add 5 ml of filter sterilized 40% urea solution. Mix, distribute in 2–4 ml amounts in 12×100 mm test tubes. Allow the medium to solidify in a slanting position in such a way to get half inch butt and one inch slant.

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Urease Test

Table 23-1 Urease producing bacteria and fungi Urease producing bacteria Strong (or) most rapid urease producers Brucella species Helicobacter pylori Rapid urease producers Proteus species Morganella species Slow urease producers Klebsiella species Enterobacter species Urease producing fungi

NEGATIVE

POSITIVE

Cryptococcus neoformans Trichophyton mentagrophytes FIGURE 23-1 Urease negative and positive test.

QUALITY CONTROL Positive control: P. mirabilis (urease positive bacteria). Negative control: Escherichia coli (urease negative bacteria). An uninoculated medium is incubated along with the test to compare the colour change.

OBSERVATION Examine the medium after four hours and after overnight incubation.The test should not be considered negative till after four days of incubation.

The uninoculated medium is colour less. In a positive test, after incubation, the colour of the medium changes to purple pink.

RESULTS AND INTERPRETATION Positive reaction is detected after 18 hours of incubation. When positive, the color of the medium changes to purple pink (P. mirabilis).The test is considered negative if no colour change of the medium is observed (E. coli) (Fig. 23-1). P. mirabilis tested is a urease producing bacteria. E. coli does not produce the enzyme urease.

KEY FACTS 1 Certain bacteria and fungi possess the enzyme urease that hydrolyzes urea releasing ammonia into the medium. 2 Phenol red that is incorporated in the medium changes its color from yellow to red in alkaline pH, thus indicating the presence of urease activity. 3 Control strains should be used for interpretation of results.

VIVA 1. Describe principle of the urease test. 2. List different media used for testing urease activity of the microorganism and how do you interpret the results? Ans. Christensen’s urea agar is a solid medium whereas Stuart’s broth is a liquid medium used for testing the urease activity of the bacteria. In Christensen’s agar, purple pink colour is seen throughout the medium which indicates alkalinization and urea hydrolysis. In Stuart’s broth, purple pink colour throughout the medium indicates alkalinization and urea hydrolysis. 3. List the compositions of the Christensen’s urea agar medium. 4. Name the slow, rapid, and most rapid urease producing bacteria. 5. List urease producing microorganisms. 6. List an important use of urease test in mycology. Ans: Urease test is used in mycology for identification of urease positive Cryptococcus neoformans.

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FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

24

Indole Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine the ability of bacteria to degrade the amino acid tryptophan. 2 Distinguish the bacteria based on the indole activity.

INTRODUCTION

Kovac’s reagent consists of para-dimethyl amino benzaldehyde, 5.0 gm; isoamyl alcohol, 75.0 ml; and concentrated hydrochloric acid, 25.0 ml. Ehrlich’s reagent consists of p-dimethyl amino benzaldehyde, 4.0 gm; absolute ethyl alcohol, 380.0 ml; and concentrated hydrochloric acid, 80.0 ml. III Specimen 24 hours to 48 hours peptone water culture of Escherichia coli incubated at 37°C.

Tryptophan is an essential amino acid that can undergo oxidation by enzymatic activities of some bacteria. Conversion of tryptophan into metabolic products is mediated by the enzyme tryptophanase. The metabolic end products are indole, skatole and indole acetic acid. The ability to hydrolyse tryptophan with the production of indole is not a characteristic of all bacteria.Only some bacteria produce indole.

1 Take 0.5 ml of 24 hours to 48 hours peptone water cultures of E. coli in a small test tube. 2 Add 0.2 ml of Kovac’s reagent to the peptone water and shake. 3 Allow it to stand for few minutes and read the result.

PRINCIPLE

QUALITY CONTROL

Tryptophan is an amino acid.This is present in peptone water of the culture medium, and when acted upon by the enzyme tryptophanase, it is converted into indole, skatole and indole acetic acid. The indole reacts with aldehydes to produce a red coloured product. The aldehyde used in the test is para dimethyl amino benzaldehyde. List of indole positive and negative bacteria are presented in the Table 24-1

Positive control: E. coli (indole positive bacteria). Negative control: Klebsiella pneumoniae (indole negative bacteria).

PROCEDURE

OBSERVATION In a positive test, a red-violet ring develops within minutes on addition of Kovac’s reagent. In a negative test a yellow ring appears.

REQUIREMENTS RESULTS AND INTERPRETATION I Equipments Incubator. II Reagents and lab wares Peptone water / tryptone broth, Kovac’s reagent, or Ehrlich’s reagent, glass tubes and inoculating wire.

Positive indole test is indicated by the appearance of red-violet ring on adding the reagent. Negative reaction is indicated by developing a yellow ring (Fig. 24-1). E. coli colonies tested are an indole producing bacteria. K.pneumoniae does not produce the indole

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Table 24-1 List of indole positive and negative bacteria Indole positive bacteria

Indole negative bacteria

1. Escherichia coli 2. Klebsiella oxytoca 3. Proteus vulgaris 4. Morganella morganii 5. Providencia rettgeri 6. Aeromonas hydrophila 7. Pasteurella multocida 8. Vibrio cholerae 9. Falvobacterium 10. Plesiomonas shigelloides

1. Escherichia vulnaris 2. Klebsiella pneumoniae 3. Proteus mirabilis 4. Salmonella Typhi 5. Shigella sonnei

NEGATIVE

POSITIVE

FIGURE 24-1 Indole negative and positive test.

KEY FACTS 1 End product of tryptophan metabolism by tryptophanase is indole, skatole, and indole acetic acid. 2 Indole reacts with p-dimethyl amino benzaldehyde to form Quinoidal red-violet compound.

VIVA 1 2 3 4 5 6

What are the end products of tryptophan metabolism? What is the principle behind the indole production test? What are the reagents used in this test? What are the constituents of Kovac’s reagent? What are the constituents of Ehrlich’s reagent? How to test indole production by adding Ehrlich reagent. Ans.Take 0.5 ml of culture broth and mix with equal volumes of Ehrlich’s reagent.Shake and allow it to stand for few minutes then observe the result. Positive reaction is indicated by development of red-violet ring and negative by yellow ring. This can be also tested in another method by adding 0.5-1 ml of xylene or ether to the culture broth and shaking it well. Then 0.5 ml of Ehrlich’s reagent is added and the result read.

FURTHER READINGS

1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

25

Methyl Red Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine the ability of bacteria to oxidise glucose with the production of high concentrations of acidic end products by methyl red test. 2 Differentiate between all glucose oxidizing enteric bacteria particularly Escherichia coli and Enterobacter aerogenes.

INTRODUCTION All enteric bacteria ferment glucose with the production of organic acids and energy.The hexose monosaccharide glucose is the major substrate oxidized by the enteric bacteria. The end products of this process vary depending on the specific enzymatic pathways present in the bacteria. In MR test the methyl red is the pH indicator. The methyl red detects the presence of large concentrations of acidic end products.This test is of value to differentiate between E. coli and E. aerogenes, and other members of the family Enterobacteriacae. Both these bacteria initially produce organic acidic products during the early period of incubation. The low acidic pH (4) is stabilized and maintained by E. coli at the end of incubation. During the later period of incubation, E. aerogenes enzymatically converts these acids to non-acidic end products such as 2, 3 – butanediol and acetoin (acetyl methyl carbinol) resulting in an elevated pH of 6.

The methyl red test is a quantitative test for acid production, requiring positive organisms to produce strong acids (lactic acid, acetic acid, formic acid) from glucose through the mixed acid fermentation pathway. Because many species of the Enterobacteriaceae may produce sufficient quantities of strong acids that can be detected by methyl red indicator during the initial phase of incubation, only organisms that can maintain this low pH after prolonged incubation (48–72 hours) overcoming the pH buffering system of the medium can be called methyl red positive. List of MR positive and negative bacteria is presented in the table 25-1.

REQUIREMENTS I Equipments Incubator. II Reagents and lab wares Inoculating wire. Methyl red test broth. It consists of poly peptone, 7 gm; glucose, 5 gm; dipotassium phosphate, 5 gm; and distilled water, 1l at a pH of 6.9. Methyl red indicator. It consists of methyl red, 0.1 g in 300 ml of 95% ethyl alcohol. III Specimen Culture of E. coli, E. aerogenes and Klebsiella pneumoniae in glucose phosphate medium incubated at 30°C for five days.

PRINCIPLE PROCEDURE Methyl red is a pH indicator with a range between 6.0 (yellow) and 4.4 (red). The pH at which methyl red detects acid is considerably lower than the pH of other indicators used in bacteriologic culture media. Thus to produce a colour change, the test organism must produce large quantities of acid from the carbohydrate substrate being used.

1 Take 0.5 ml of broth cultures of E. coli in a small test tube. 2 Add five drops of 0.04% solution of methyl red directly to the broth culture and mix well. 3 Note any change in the colour of medium at once.

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Table25-1 List of MR positive and negative bacteria

RESULTS AND INTERPRETATION

MR positive bacteria

MR negative bacteria

1. E. coli 2. K. ozaenae 3. K. rhinoscleromatis 4. K. ornitholytica 5. Edwardsielleae 6. Salmonellae 7. Citrobacter 8. Proteae 9. Yersinia

1. K. pneumoniae 2. Enterobacter spp

The development of a stable red colour in the surface of the medium indicates sufficient acid production to lower the pH to 4.4 and constitutes a positive test. Because other organisms may produce smaller quantities of acid from the test substrate, an intermediate orange colour between yellow and red may develop. This does not indicate a positive test. Yellow colour indicates a negative test (Fig. 25-1).

QUALITY CONTROL Positive control: E. coli Negative control: E. aerogenes

OBSERVATION Look for the development of stable red colour on adding methyl red indicator.

INDOLE

MR

VP

CITRATE

FIGURE 25-1 IMViC Test.

KEY FACTS 1 2 3 4 5

MR test detects the products of stable high concentration of acidic end products. The test differentiates between E. coli and E. aerogenes. Methyl red is the indicator used in the test. Methyl red is yellow in alkaline pH and red in acidic pH. Development of red colour on addition of methyl red to broth culture of bacteria is considered positive.

VIVA 1 2 3 4

What are the group of organisms that can be differentiated by MR test? What is the principle of MR Test? What are positive and negative controls used in the test? How long you have to incubate the test and why? Ans: The broth is incubated for 48-72 hr, because the organisms have to produce sufficient quantities of acid and it has to be maintained in the same acidic pH which turns the methyl red indicator to red colour. 5 What is the indicator used in this test?

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

26

Voges-Proskauer Test

LEARNING OBJECTIVES After completing this practical you will be able to:

small amount of acetyl methyl carbinol present in the medium is converted to diacetyl, which reacts with the peptone of the broth to produce a red colour. List of VP positive and negative bacteria is presented in the table 26-1.

1 Perform Voges-Proskauer test.

REQUIREMENTS INTRODUCTION Voges-Proskauer is a double eponym, named after two microbiologists working at the beginning of the 20th century. They first observed the red colour reaction produced by appropriate culture media after treatment with potassium hydroxide. It was later discovered that the active product in the medium formed by bacterial metabolism is acetyl methyl carbinol, a product of the butylene glycol pathway.

PRINCIPLE The Voges-Proskauer test determines the capability of some bacteria to produce non-acidic or neutral end products such as acetyl methyl carbinol from the organic acids produced as a result of glucose metabolism. Pyruvic acid, the pivotal compound formed in the fermentative degradation of glucose is further metabolised through various metabolic pathways, depending on the enzyme systems possessed by different bacteria. One such pathway results in the production of acetoin (acetyl methyl carbinol), a neutral-reacting end product. Enteric bacteria such as members of the Klebsiella-EnterobacterHafnia-Serratia group produce acetoin as the chief end products of glucose metabolism and form smaller quantities of mixed acids. The test depends on the production of acetyl methyl carbinol from pyruvic acid, as an intermediate product in its conversion to 2: 3 butylene glycol. In the presence of atmospheric oxygen and alkali (40% potassium hydroxide), the

I Equipments Incubator. II Reagents and lab wares Inoculating loop. VP broth. It consists of polypeptone, 7 gm; glucose,5 gm; dipotassium phosphate, 5 gm and distilled water, 1 litre at a pH of 6.9. 5% a naphthol. It consists of a naphthol, 5 gm; and absolute ethyl alcohol, 100 ml. It serves as the colour intensifier. 40% potassium hydroxide. It consists of 40 gm potassium hydroxide in 100 ml distilled water. It serves as the oxidising agent. III Specimen Culture of Escherichia coli, Enterobacter aerogenes and Klebsiella pneumoniae in glucose phosphate medium incubated at 30°C for five days or 37°C for 48 hours.

PROCEDURE 1 Take 1 ml of broth cultures of E. coli in a small test tube. 2 First add 40% KOH and then add 0.6 ml of a 5% solution of a naphthol in ethanol to the broth culture and shake gently. It is essential that the reagents are added in this order. 3 Note any change in the colour of medium within 2-5 minutes.

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QUALITY CONTROL Positive control: E. aerogenes. Negative control: E. coli.

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hour.The test should not be read after standing for over 1 hour because negative VP test may produce a copper-like colour, leading to a false positive interpretation.

Table 26-1 VP positive and negative bacteria

OBSERVATIONS Look for the development of pink colour 15 minutes or more after addition of the reagents.

RESULTS AND INTERPRETATION A positive test is represented by the development of a pink colour 15 minutes or more after addition of the reagents, deepening to magenta or crimson in half an hour.This indicates the presence of diacetyl, the oxidation product of acetoin. A negative test is indicated by colour less reaction for half an

VP positive bacteria

VP negative bacteria

1. Klebsiella pneumoniae 2. Enterobacter cloacae 3. Cedicia netri 4. Ewingella americana 5. Serratia marcescens 6. Aeromonas sobria 7. Vibrio cholerae 8. Chryseomonas luteola 9. Flavimonas oryzihabitans 10. Sphingomonas paucinobilix

1. Escherichia coli 2. Edwardsiella tarda 3. Salmonellae 4. Proteae 5. Yersinieae

KEY FACTS 1 The VP test shows the presence of non acidic neutral end product acetyl methyl carbinol. 2 A positive test is represented by the development of a pink colour 15 minutes or more after addition of the reagents, deepening to magenta or crimson in half an hour. 3 A negative test is indicated by colour less reaction for half an hour. 4 Some times prolonged incubation of cultures is required before doing the test.

VIVA 1 2 3 4

What is the principle of VP test? What are the positive and negative controls used in the VP test? Name some VP positive bacteria? Describe the biochemical reaction in the VP test.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

27

Citrate Utilisation Test

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to: 1 Differentiate certain enteric organisms on the basis of their ability to utilize citrate as a sole source of carbon.

I Equipments Incubator.

INTRODUCTION Sodium citrate is a salt of citric acid, a simple organic compound produced as one of the metabolites in the tricarboxylic acid cycle of the bacteria. Some bacteria can also obtain energy by using citrate as the sole source of carbon. Hence, any medium used to demonstrate citrate utilisation by test bacteria must be devoid of protein and carbohydrates as sources of carbon.

II Reagents and lab wares Inoculating loop, Simmon’s citrate medium (It consists of ammonium dihydrogen phosphate, 1 gm; dipotassium phosphate, 1 gm; sodium chloride, 5 gm; sodium citrate, 2 gm; magnesium sulfate, 0.20 gm; agar, 15 g; bromo thymol blue, 0.08 gm and distilled water 1 litre) pH adjusted to 6.9. The medium is poured into a tube on a slant. III Specimen Culture of Escherichia coli, Enterobacter aerogenes and Klebsiella pneumoniae in glucose phosphate medium incubated at 37°C for 48 hours.

PRINCIPLE In the absence of fermentable glucose or lactose, some bacteria are capable of using citrate as a sole source of carbon for their energy. This ability depends on the presence of the enzyme, a citrate permease that facilitates the transport of citrate in the cell. Citrate is the first major intermediate in the Krebs cycle and is produced by the condensation of active acetyl with oxaloacetic acid. Citrate is acted on by the enzyme citrase which produces oxaloacetic acid and acetate. These products are then enzymatically converted to pyruvic acid and carbon dioxide (CO2). During this reaction the medium becomes alkaline because the CO2 that is generated combines with sodium and water to form sodium carbonate, an alkaline product. The presence of sodium carbonate changes the indicator, bromo thymol blue present in the medium from green at pH 6.9 to deep Prussian blue at pH 7.6. Simmon’s citrate and Koser’s citrate are two examples of different types of citrate media used in the test. Differences between the two media are summarized in the table 27-1. List of citrate positive and negative bacteria are presented in the table 27-2.

PROCEDURE 1 Using sterile technique, inoculate each bacteria into its appropriately labeled tube by means of a stab and streak inoculation. 2 Incubate all cultures for 24 hours to 48 hours at 37°C. Table 27-1 Differences between Simmon’s citrate and Koser’s citrate Simmon’s citrate

Koser’s citrate

It is a slant (solid medium) It contains agar This medium contains bromothymol blue as indicator Positive test is indicated by growth on the medium and change in the colour of the medium.

It is a broth (liquid medium) It contains no agar It does not contain any indicator Positive test is by observing the turbidity in the medium

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QUALITY CONTROL

RESULTS AND INTERPRETATION

Positive control: E. aerogenes. Negative control: E. coli.

A positive test is represented by the development of a deep blue colour within 24 hours to 48 hours, indicating that the test organism has been able to utilize the citrate contained in the medium, with the production of alkaline products. A negative test is indicated by no change of colour of the citrate medium (Fig. 27-1).

OBSERVATIONS Look for the development of deep blue colour within 24-48 hours of incubation of the inoculated tube.

Table 27-2 List of citrate positive and negative bacteria Citrate positive bacteria

Citrate negative bacteria

1. Klebsiella pneumoniae 1. Escherichia coli 2. Citrobacter diversus 2. Salmonella Typhi 3. Enterobacter cloacae 3. Salmonella Paratyphi A 4. Serratia marcescens 4. Shigella species 5. Providencia alcalifaecians 5. Yersinia enterocolitica 6. Euringella americana 6. Edwardsiella tarda 7. Acroncobacter oxylosoxidans 7. Vibrio holisae 8. Vibrio vulnificus

NEGATIVE

POSITIVE

FIGURE 27-1 Citrate negative and positive test.

KEY FACTS 1 2 3 4

Simmon’s citrate medium contains sodium citrate which acts as a sole source of carbon. Bromothymol blue is the indicator used in Simmon’s citrate medium. Development of deep blue colour is taken as positive. A negative test is indicated by no change of colour of the citrate medium.

VIVA 1 2 3 4 5

What are the constituents of Simmon’s citrate medium? What is the principle behind citrate utilisation test? What is the positive control and negative controls used in the citrate utilisation test? What are the organisms that are citrate utilisation positive and negative? How will you interpret the results of citrate utilisation test? Ans: A positive test is represented by the development of a deep blue colour within 24 hours to 48 hours indicating that the test organisms has been able to utilize citrate contained in the medium, with the production of alkaline products. 6 Mention different types of citrate utilization tests. Ans: Simmon’s citrate utilization test and Koser’s citrate utilization test. 7 List differences between Koser’s citrate and Simmon’s citrate media.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

28

Triple Sugar Iron (TSI) Agar Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Differentiate among the members of the family Enterobacteriaceae by the TSI test.

INTRODUCTION The triple sugar iron (TSI) agar is an example of a composite medium used widely for the identification of bacterial isolates.This medium is convenient and economical, because as a single composite medium different properties of the bacteria which otherwise would have required the use of many separate media could be used. The TSI agar is designed to differentiate among different groups or genera of the family Enterobacteriaceae. The latter are the Gram Negative bacilli capable of fermenting glucose with the production of acid. The differentiation can be made on the basis of differences in carbohydrate fermentation and hydrogen sulfide production by various intestinal bacteria. The TSI agar has glucose, lactose, and sucrose as the sources of carbohydrates. The slant contains lactose and sucrose in the concentration of 1% and glucose in the concentration of 0.1%.Phenol red is the acid base indicator incorporated in the medium.This indicator helps to detect carbohydrate fermentation that is indicated by a change in colour of the medium from orange red to yellow in the presence of acid.

PRINCIPLE The TSI agar is distributed in the tube which contains a slant and a butt.TSI medium indicates whether the bacteria ferments glucose only, or lactose and sucrose also with or without production of gas. The medium can detect production of hydrogen sulphide (H2S) as well as other bacteria which utilizes only glucose (but not lactose or sucrose). Due to the acid production, the colour of the phenol red (indicator) is changed

to yellow and the whole medium appears yellow in colour. After further incubation as the glucose is fully exhausted, the bacteria begin to oxidatively degrade the amino acid present in the medium. Since oxygen is exposed only to the slant portion, oxidative degradation occurs only in the slant portion. This oxidative degradation results in production of alkali products, which reverts the colour of the slant to red colour. In the deeper part of the tube, amino acid degradation is insufficient to overcome the acid formed, so medium in the butt part remains yellow in colour. If the TSI medium is inoculated with lactose fermenting organism, then even after the glucose is completely used up in first 8–12 hours, fermentation continues as the organism is able to use lactose which is present in concentration 10 times that of glucose. So the acid production continues to occur even after 18–24 hours and both the slant and the butt appear yellow. For the detection of H2S which is a colourless gas, medium must include an indicator to detect the H2 S. Sodium thiosulfate is the source of sulfur atoms. Ferrous sulfate is the indicator used for the detection of the H2S which is indicated by the production of insoluble black precipitate.

REQUIREMENTS I Equipments Incubator. II Reagents and lab wares Inoculating loop and triple sugar iron agar (It contains beef extract, 3g; yeast extract, 3g; peptone, 15g; proteose peptone, 5g; lactose, 10g; sucrose, 10 gm; glucose, 1 gm; ferrous sulfate, 0.2 gm; sodium chloride, 5g; sodium thiosulfate, 0.3g; agar, 12 gm; phenol red, 0.024 g; and distilled water to equal 1 litre) at pH 7.4. III Specimen Culture of test organisms such as Escherichia coli, Proteus mirabilis and Klebsiella pneumoniae in glucose phosphate medium incubated at 37°C for 48 hours.

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PROCEDURE 1 Using sterile technique, inoculate each bacterial colony into its appropriately labeled tube by means of a stab and streak inoculation. 2 Incubate all cultures for 18 hours to 24 hours at 37°C.

QUALITY CONTROL 1 Alkaline slant /alkaline butt (K/K reaction): Pseudomonas aeruginosa. 2 Alkaline slant /acidic butt (K/A reaction): Shigella, Vibrio. 3 Alkaline slant /acidic butt /production of H2 S (K/A reaction and positive for H2 S): Salmonella spp, Citrobacter spp, Proteus spp. 4 Acidic slant /acidic butt (A/A reaction): E. coli, Klebsiella spp, Enterobacter spp. 5 Acid butt/acid slant, H2 S positive: Citrobacter.

OBSERVATIONS Look for the colour change in the slant and butt after 18–24 hours incubation and also look for the development of black precipitate to indicate H2S production (Fig.28-1).

RESULTS AND INTERPRETATION 1 Alkaline slant /alkaline butt (K/K reaction): This shows no carbohydrate fermentation. This indicates that the bacteria are non-fermented. Example: P. aeruginosa.

A+/A-

Un inoculated

K-/K-

K+/A+

FIGURE 28-1 TSI reactions.

2 Alkaline slant /acidic butt (K/A reaction): Glucose is fermented. Lactose and sucrose is not fermented. It indicates that the organism is a non -lactose fermenter. Example: Shigella, Vibrio. 3 Alkaline slant /acidic butt /black precipitate of H2S (K/A reaction and positive for H2S): Glucose is fermented. Lactose and sucrose are not fermented. This is characteristic of non -lactose fermenting, hydrogen sulfide producing bacteria. Example: Salmonella spp, Citrobacter spp, Proteus spp. 4 Acidic slant /acidic butt (A/A reaction): All the sugars (glucose, lactose, and sucrose) are fermented. This is characteristic of lactose fermenting coliforms. Example E. coli, Klebsiella spp, Enterobacter spp.

VIVA 1. What are the constituents of TSI medium? 2. What is the source of sulfur? 3. What is the source of nutrition? Ans: Sucrose, lactose, glucose and peptone 4. What are the sugars used and at what concentration they are used in the TSI agar? 5. What is the principle behind this test? 6. What is the H2S indicator? 7. When the reading of TSI test is taken and why? Ans: The readings of TSI test is taken only after 18-24 hrs. The reason behind this is that the glucose concentration is 10 times less than that of lactose and sucrose. Glucose fermenters break down glucose first then lactose. If the organism is non lactose or sucrose fermenter, the organisms metabolise peptones aerobically for their energy, producing alkaline end products. In the first 6-8 hours whole of the tube will be acidic and will be yellow in colour. Production of alkaline end products results in the change of pH alkaline side and pink colour develops in the slant. So the pH of the slant is alkaline after 8 hours. So the results are read after 24 hours.

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Triple Sugar Iron (TSI) Agar Test

KEY FACTS 1 2 3 4 5 6

Beef extract, yeast extract, peptone, and proteose peptone makes the medium nutritionally rich. Glucose and lactose; and sucrose are added in the ratio of 1: 10 in the TSI agar. Oxidative degradation of amino acids in the medium occurs only in the slant. Ferrous sulfate is the indicator for the production of the hydrogen sulfide. Sodium thiosulfate is the source of sulfur. Phenol red is the indicator used to detect acid /alkaline changes in the medium.

FURTHER READINGS

1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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LESSON

29

Hydrogen Sulfide Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine the ability of certain bacteria to produce hydrogen sulfide from substrates such as the sulfur containing amino acids or inorganic sulfur compounds.

INTRODUCTION Some bacteria liberate sulfur from sulfur containing amino acids or other sulfur containing compounds. The sulfur is used as final hydrogen acceptor leading to the formation of hydrogen sulphide (H2S). Sulfur containing amino acids such as methionine, cystine, cysteine or inorganic compounds such as this sulphates, etc. should be present in the medium to detect the presence of the H2 S. H2S being a gas, will escape from the medium. So, the indicators such as heavy metal ions are added to the medium, which support the growth of the organism. Bacteria which are tested for the production of H2 S must contain enzyme system which release sulfide from the sulfur source. Sulfides combine with hydrogen ion to form H2 S. H2 S combines with heavy metals to form insoluble black precipitate.

PRINCIPLE There are two major fermentative pathways by which hydrogen sulfide is produced by bacteria. Pathway 1: Gaseous H2S may be produced by the reduction (hydrogenation) of organic sulfur present in the amino acid cysteine, which is the source of sulfur in the medium. This amino acid in the presence of the enzyme, a cysteine desulfurase, loses the sulfur atom and is then reduced by the addition of hydrogen from water to form hydrogen gas. Pathway 2: Gaseous H2S may also be produced by the

reduction of inorganic sulfur compounds such as the thiosulfates (S2O3), sulfates (SO4) or sulfites (SO3). The medium contains sodium thiosulfate, which are reduced to sulfite by certain microorganisms with the liberation of hydrogen sulfide. The sulfur atoms act as hydrogen acceptors during oxidation of the inorganic compound. Liberated H2S combines with indicator like ferrous sulfate (Fe SO4) to produce black insoluble precipitate (ferrous sulfide). List of media used for detecting production of hydrogen sulphide is presented in the box 29-1. List of bacteria producing hydrogen sulphide is presented in the table 29-1.

REQUIREMENTS I Equipments Incubator. II Reagents and lab wares Sulphur containing medium such as Kligler iron agar, TSI agar or lead acetate agar, etc. and inoculating loop. III Specimen Soy broth cultures of test bacteria such as Enterobacter aerogenes, Proteus vulgaris and Salmonella Typhimurium incubated at 37°C for 24- 48 hours.

PROCEDURE 1 Using sterile technique, pick up the organisms from the top of a single colony from primary isolation plate or from pure growth with a straight wire. 2 Inoculate by stabbing down the center of agar butt carefully and then streak the surface of the slant. 3 Incubate all cultures for 18 hours to 24 hours at 37°C.

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BOX 29-1 LIST OF MEDIA USED FOR DETECTING PRODUCTION OF HYDROGEN SULFIDE List of media 1. Bismuth sulfite agar 2. Citrate sulfide agar 3. Deoxycholate citrate agar 4. Lysine iron agar. 5. Kligler iron agar 6. TSI agar 7. Lead acetate agar 8. Salmonella-Shigella agar 9. Sulphur indole motility medium 10. Xylose lysine deoxycholate agar 11. Hektoen enteric agar 12. Peptone water, lead acetate paper inserts. Note: Source of sulfur and H2S indicators are different for each medium. Sources of sulfur 1. Sulfite 2. Peptone 3. Sodium thiosulfate

Table 29-1 List of H2S positive bacteria 1 Citrobacter freundii 2 Salmonella Arizona 3 Salmonellae spp (except S. Paratyphi A) 4 Proteus mirabilis 5 Proteus vulgaris 6 Edwardsiella tarda 7 Edwardsiella hoshinae 8 Shewanella putrefaciens 9 Campylobacter sputorum 10 Brucella abortus 11 Brucella suis. 12 Erysiphilothrix rhusiopathiae

RESULTS AND INTERPRETATION Black coloration along the streak line or throughout the medium indicates H2S production. If black colour is not produced then H2S is not produced (Fig. 29-1).

H2S indicator 1. Ferrous sulfate 2. Ferric ammonium citrate 3. Lead acetate 4. Peptonised iron.

QUALITY CONTROL Positive control: P. vulgaris, S. Typhimurium. Negative control: E. aerogenes.

OBSERVATIONS Look for the development of black coloured streak line after 18–24 hours incubation.

NEGATIVE

FIGURE 29-1 TSI agar showing absence and presence of H2S.

VIVA 1 2 3 4 5 6 7 8

POSITIVE

What are the conditions required for the production and demonstration of H2S by bacteria? What are the sequence of events occurring in the production and detection of H2S? What is the source of sulfur in the media? What are the indicators of H2S production? What are the sulfur containing amino acids? How will you pick up the colonies for inoculation? What are the media that can be used in H2S production detection? Name some H2S producing bacteria.

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KEY FACTS 1 Requirements to demonstrate H2S production. • A source of sulfur in the medium. • A H2S indicator in the medium. • Medium must support the growth of bacteria being tested. • The bacteria must possess the enzyme system to produce H2S. 2 First step in the production of H2S is release of sulfide ions from sulfur containing amino acids or thiosulfates by bacterial enzymes. 3 Second step is the coupling of sulfide to H+ ions to form H2S. 4 Third step is H2S reacting with heavy metals like iron, bismuth, lead to produce insoluble heavy metal sulfides that appear black precipitate. 5 Cysteine, cystine, methionine are some sulfur containing amino acids. 6 Only tops of the colonies growing on selective media must be touched, since inhibited flora may still be present and viable. These may interfere with the reaction. Otherwise single colony of purified culture must be used. 7 Sources of sulfur in the medium may be sulfite, sodium thiosulfate, or peptone. 8 Different H2S indicators used in various media are ferrous sulfate, ferric ammonium citrate, lead acetate and peptonised iron. FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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30

Nitrate Reduction Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine the ability of certain bacteria to reduce nitrates (NO3–) to nitrites (NO2–) or beyond the nitrite stage.

INTRODUCTION Nitrates serve as a source of nitrogen for many bacteria. They can also act as final electron acceptor. Many bacteria can be differentiated and are identified by their capacity to reduce nitrates to nitrites. Most of the bacteria belonging to the family Enterobacteriaceae reduce nitrates. This character is also useful for the identification of species in the genera Neisseria, Haemophilus and Branhamella. Some Pseudomonas and nonfermenters reduce nitrate to nitrite and further down to N2 and molecular nitrogen. This is called denitrification.

extract, 3 gm; peptone,5 gm; potassium nitrate,1 gm; agar,12 gm, and distilled water, 1 L). Reagent A (a naphthylamine, 5 g; acetic acid (5N) 30%, 1 L). Reagent B (sulphanilic acid, 8 g; acetic acid (5N) 30%, 1 L). Inoculating loop. III Specimen Cultures of test bacteria such as Escherichia coli, Acinetobacter baumannii, etc. in a broth containing 1% potassium nitrate broth (KNO3) incubated at 37°C for 5 days.

PROCEDURE 1 Mix an equal volume (0.5 ml) of reagent A with 0.5 ml of reagent B just before use. 2 Add 0.1 ml of the test reagent to 1 ml of the culture broth of the bacteria to be tested. 3 Observe for any change of colour immediately within few minutes.

PRINCIPLE Bacteria demonstrating nitrate reduction have the capability of extracting oxygen from nitrates to form nitrite and other reduction products. The chemical reaction is NO3–+ 2e– + 2H+®NO2– + H2O. The presence of nitrites in the test medium is detected by the addition of a-naphthylamine and sulphanilic acid, with the formation of a red diagonium dye, p-sulfobenzena-azo-anaphthylamine.

QUALITY CONTROL Positive control: E. coli. Negative control: A. baumannii.

OBSERVATION

REQUIREMENTS

Observe for the development of red colour immediately within few minutes of adding the reagents.

I Equipments Incubator.

RESULTS AND INTERPRETATION

II Reagents and lab wares 1% potassium nitrate broth (KNO3) or nitrate agar slant (beef

The development of a red colour within 30 seconds. after adding the test reagents indicates the presence of nitrites and represents positive reaction (Fig. 30-1).

Textbook of Practical Microbiology

If no colour develops after adding the test reagents, it is taken as negative test. It might have been reduced to products other than nitrites such as molecular nitrogen, nitric oxide or nitrous oxide. In this case, the reaction may show a false negative reading. Thus it is necessary to add a small quantity of zinc dust to all negative reactions. Zinc dusts reduce nitrates to nitrites, and the development of a red colour after adding zinc dust indicates the presence of residual nitrates and confirms a true negative reaction.

NEGATIVE

89

POSITIVE

FIGURE 30-1 Nitrate reduction test.

KEY FACTS 1 Nitrates are reduced to nitrites by some organisms. 2 Two reagents used in the test are: a naphthylamine and sulphanilic acid. 3 Zinc dust is added to detect all negative reactions.

VIVA 1 2 3 4 5

What is the principle behind nitrate reduction test? What are the ingredients present in nitrate broth? What are reagents added in the reactions? What is the positive and negative control for nitrate reduction test? What is the purpose of adding zinc dust to all negative reaction? Ans: Negative reaction in nitrate reduction test may be of 2 types. One is the true negative reaction in which nitrates are not reduced to nitrites or in false negative reactions the nitrates are reduced to products other than nitrites such as N2, NO or N02. The reagents can detect only the presence of nitrites. Zinc ions reduces nitrates to nitrites and the development of a red colour after adding zinc dust, indicates the presence of residual nitrates and confirms a true negative reaction. 6 How will you avoid false negative reaction? Explain.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS (Eds). Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Introduction to Clinical Microbiology. University of Texas- Houston Medical School, DPALM Medic. 1995. 4 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC (Eds). Color Atlas and Textbook of Diagnostic Microbiology. 5th ed. (Lippincott Williams and Wilkins, USA). 1997.

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UNIT

V Antimicrobial Sensitivity Tests

Lesson 31 Kirby-Bauer Method Lesson 32 Stoke’s Method Lesson 33 Agar Dilution Method Lesson 34 Broth Dilution Method Lesson 35 Epsilometer Test (E-test)

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LESSON

31

Kirby–Bauer Method

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine antibacterial sensitivity of bacterial isolates by Kirby –Bauer disc diffusion method.

INTRODUCTION Due to emergence of many antibiotic resistant strains of bacteria, antimicrobial susceptibility testing is done in order to determine which antimicrobial agent to use against a specific strain of bacteria. The available chemotherapeutic agents vary in their scope of antimicrobial activity. Some have a limited spectrum while others have a wide spectrum of activities against bacteria. The bacterial strains isolated from clinical samples should be tested for antimicrobial sensitivity because it gives the clinician an idea as to what antimicrobial therapy should be started to the patients (Box 31-1).

PRINCIPLE Kirby – Bauer method is a method of determination of antibiotic sensitivity of the bacteria by disc diffusion method. In this method, a standard suspension of bacteria to be tested are inoculated on the surface of Mueller Hinton agar plates. Filter paper discs containing specific concentration of antimicrobial agents are pressed on to the surface and incubated at 35°C overnight (18-24 hr.). After incubation, the zone of inhibition of growth of bacteria around each disc is measured and the susceptibility is determined.

REQUIREMENTS I Equipments Incubator.

II Reagents and lab wares 0.5 McFarland standard, Mueller Hinton agar plates (pH 7.27.4), peptone water , filter paper discs impregnated with appropriate concentration of antibiotics, sterile cotton swabs, millimeter ruler, forceps and inoculating wire. Preparation of 0.5 McFarland standard: Solution A is prepared by adding barium chloride (BaCl 2, 2H2O) to 100ml distilled water. Solution B is prepared by adding 1ml of sulphuric acid (H2S04 (0.36N) to 100 ml of distilled water. Then 0.5 ml of solution A is added to 99.5 ml of solution B, mixed well and distributed in test tubes with a screw cap. The cap is closed tightly to avoid evaporation. The mixture is stored in the dark. The solution is agitated vigorously before using it. III Specimens Staphylococcus aureus ATCC 25923, Escherichia coli ATCC 25922, Enterococcus faecalis ATCC 29212, and Pseudomonas aeruginosa ATCC 27853. Preparation of suspension of bacteria: Approximately, 4-5 well isolated colonies of the bacterial strain to be tested are inoculated into 5 ml of peptone water, and is incubated at 37 °C for 3-4 hours. The turbidity of the suspension is adjusted to match 0.5 McFarland standards. If the density is more it is diluted with sterile saline. The comparison is made against a white back ground with a contrasting black line.

PROCEDURE 1 After standardisation of bacterial suspension, immerse a sterile cotton swab in it and rotate the swab several times with firm pressure on the inside wall of the tube to remove excess fluid. 2 Prepare a Mueller Hinton agar (MHA) plate (pH 7.2-7.4) with a depth of 4 mm. 3 Inoculate the dried surface of the MHA agar plate by streaking the swab three times over the entire agar surface. It is streaked in three directions by rotating the plate 60° after each streak.

Textbook of Practical Microbiology

4 Place the appropriate antimicrobial impregnated discs on the surface of the agar using sterile forceps. 5 Gently press each disc onto the agar to provide uniform contact. Do not move the disc once it has contacted the agar because some of the antibiotics diffuse almost immediately Discs must be placed in such a way that they are at least 20 mm from one another. Note: 6 antibiotic discs may be put in an 85 mm plate. 6 Invert the plates and incubate at 35 °C -37°C for 16-18 hr.

QUALITY CONTROL

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RESULTS AND INTERPRETATION Each antibiotics produces a specific zone size for each bacteria tested. Depending on the zone size, the bacteria are classified as follows (Fig. 31-1): Sensitive (S): Infection treatable with normal dosage of the antibiotic. Intermediate (I): Infection may respond to therapy with higher dosage Resistant (R): Unlikely to respond to the antibiotic at the usual dosage.

S. aureus ATCC 25923, E. coli ATCC 25922, E. faecalis ATCC 29212, and P. aeruginosa ATCC 27853 should be tested periodically.

OBSERVATIONS 1 Examine the plates for the presence and size of inhibitory zones. 2 The diameter of the inhibitory zone including the diameter of the disc is measured by using a millimeter scale upto the nearest millimeter. 3 All measurements are made with unaided eye while viewing the back of the petri dish with reflected light against a black non-reflecting background. 4 Measure the inhibitory zones for each antimicrobial agent, compare with the standard Kirby-Bauer’s chart and interpret the zone of inhibition as sensitive, intermediate or resistant.

FIGURE 31-1 Kirby-Baur method of antibiotic susceptibility testing.

BOX 31-1 GLOSSARY OF TERMS Antibiotics: It is a substance produced by microorganisms or a similar substance produced wholly or partially by chemical synthesis that inhibits the growth or causes death of other organisms in low concentrations. Antimicrobial agent: It is a chemical substance inhibiting the growth or causing death of microorganism. Bacteriostatic drug: Certain antimicrobial agents inhibit the growth by preventing the multiplication of organisms. They do not cause death. These are called bacteriostatic agents. eg. tetracycline, erythromycin, and sulphonamides. Bactericidal drug: The drugs that cause irreversible damage to bacteria resulting in death are called bactericidal drugs. e.g. aminoglycosides, penicillin and quinolones.

KEY FACTS 1 Depth of agar in medium should be 4 mm as some antibiotics show decreased zone size with increased depth while others show slight increase. 2 Standardization of the bacterial inoculum is important. It should be such that it gives rise to a semi confluent growth, as growth denser than this or lighter than this give problem while reading zone size. 3 Proper storage of the antimicrobial discs, so that they retain their potency.

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VIVA 1 Define a) antibiotics, b) antimicrobial agent, c) bacteriostatic drug and d) bactericidal drug. 2 What is the importance of antimicrobial susceptibility testing? 3 Why is the standardization of pH of medium important in disc diffusion test? Ans: Standardization of pH of the medium is important because some antibiotics show increased or decreased zone sizes depending on pH. Antibiotics stimulated by fall in pH i.e. larger zone sizes – Tetracycline, methicillin, and novobiocin. Antibiotics stimulated by an alkaline pH – Aminoglycosides and macrolides such as erythromycin. 4 How are antimicrobial discs prepared? Ans: 1 Discs are made from Whatman filter paper No. 1 using standard stationary paper punch. 2 Discs are arranged separately in a Petri dish and sterilized in hot air oven at 160°C for 1 hour. 3 A 27 SWG needle, bevel cut off is used to drop the antibiotics. It is attached to a tuberculin syringe fitted with a teat. This delivers 140 drops/ ml. 4 A 6 mm disc is able to completely absorb the fluid volume (7 µl). 5 Each disc is impregnated with a drop of the solution and allowed to dry in the incubator at 37°C for 30 mins. 6 Any disc that is not to be used on the day of preparation is stored at -20 °C to 8 °C.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. 1997. pp. 1395. Lippincott Williams and Wilkins. 4 Lalitha M.K. Manual on Antimicrobial Susceptibility Testing. Christian Medical College, Vellore, 2004, pp 43. 5 WHO. Guidelines on Standard Operating Procedures for Microbiology. Blood safety and Clinical Technology. Chapter 7: Antimicrobial Susceptibility Testing.

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LESSON

32

The Stokes Method

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine antibacterial sensitivity of bacterial isolates by Stokes method.

INTRODUCTION Stokes method is an example of the disc diffusion test and is another method used for routine antibiotic sensitivity testing of bacterial strains. The method makes use of in-built controls against many variables and therefore provides dependable results. A set of standard strains are used as control strains depending on the bacterium to be tested. The control strains are Escherechia coli NCTC 10414 for testing coliform bacilli from urinary tract. Pseutomonas aeruginosa NCTC 10662 against aminoglycosides. Kirby-Bauer and Stokes methods are compared in the table 32-1.

PRINCIPLE In this test antibiotic discs are applied between the standard and test inocula, so that zones of inhibition formed around each disc are composed of standard and test bacteria. The diffusion of antibiotic takes place and thus the susceptibility of those organisms to the antibiotic are known by measuring zone size.

III Specimens Control strains: Staphylococcus aureus NCTC 6571, Escherichia coli NCTC 10414 and Pseudomonas aeruginosa NCTC 10662. Preparation of suspension of bacteria: Approximately, 45 well isolated colonies of the bacterial strain to be tested are transferred to Tryptic soy broth or BHI broth. The turbidity of the suspension is adjusted to match 0.5 McFarland standards.

PROCEDURE 1 The inoculation plates are dried with lids open so that there are no droplets of moisture on the surface. 2 The control culture is applied in two bands on either side of the plate leaving a central band uninoculated with the help of sterile swab. 3 The test organism is applied in the central portion without touching the either sides. 4 Antibiotic discs are applied with forceps on the line between the test and control organisms and pressed gently to ensure even contact with the medium. There should be a minimum distance of 2 cm between two discs. Four discs can be accommodated on an 85 mm circular plate. 5 For inoculation, a rotatory plating method can also be used wherein the control strain is applied on the outer periphery and the test strain is applied in the central portion. In such a method 6 discs can be put on an 85 mm circular plate. 6 The plates are then incubated overnight at 35°C –37°C.

REQUIREMENTS I Equipments Incubator. II Reagents and lab wares 0.5 McFarland standards, Mueller-Hinton agar plates (pH 7.27.4), antibiotics discs, sterile cotton swabs, Millimeter ruler and sterile forceps.

QUALITY CONTROL S. aureus NCTC 6571, E. coli NCTC 10414 and P. aeruginosa NCTC 10662.

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The Stoke’s Method

OBSERVATIONS Zone sizes are measured from the edge of the discs to the edge of the zone. Comparison of the zones of inhibition between the standard and test bacteria indicates the sensitivity of the test bacteria. If the test zones are obviously larger than the control or give no zone of inhibition at all; there is no need to perform any measurement with calipers or a millimeter scale.

RESULTS AND INTERPRETATION Each zone size is interpreted as follows (Fig. 32-1): Sensitive: Zone size equal to wider than or not more than 3 mm smaller than control. Intermediate: Zone size greater than 2 mm, but smaller than the control to more than 3 mm. Resistant: Zone size 2 mm or less.

FIGURE 32-1 Stokes method.

Table32-1 Comparison of Kirby-Bauer and Stokes methods Kirby-Bauer Method

Stokes Method

1. Test and control strains have to be tested on separate plates. 2. 6 discs applied on 85mm plate. 3. Zone diameter is measured including disc diameter.

1. Test and control strains tested on the same 2. 4 discs on 85mm plate and 6 discs by rotatory method. 3. Zone diameter measured from edge of disc to edge of zone of inhibitation the disc.

KEY FACTS 1 It is better to apply the control organisms followed by test organisms. 2 Penicillinase producing Staphylococci, showing heaped up, clearly defined zone edges should be reported resistant irrespective of zone size. 3 The advantage of Stoke’s method is that both control and test organisms can be tested on the same plate under similar conditions and difference between zone sizes can be directly measured.

VIVA 1 How is the zone size measured in Stokes method? 2 How many discs can be applied on a plate? Ans: A total of 4 discs can be accommodated on an 85 mm circular plate. If rotatory plating method is used 6 discs can be accommodated on an 85mm circular plate. There should be a minimum distance of 2 cm between 2 discs. 3 Compare and contrast Kirby-Bauer and Stokes methods.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. 1997. pp. 1395. Lippincott Williams and Wilkins. 4 Lalitha M.K. Manual on Antimicrobial Susceptibility Testing. Christian Medical College, Vellore, 2004, pp 43. 5 WHO. Guidelines on Standard Operating Procedures for Microbiology. Blood safety and Clinical Technology. Chapter 7: Antimicrobial Susceptibility Testing.

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LESSON

33

Agar Dilution Method

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine antibacterial sensitivity of bacterial isolates by agar dilution method.

INTRODUCTION Agar dilution is a quantitative method for determining the minimum inhibitory concentration of the antibiotics against bacteria to be tested. It is mainly useful in testing isolates from serious infections like bacterial endocarditis or to verify equivocal results (e.g. intermediate susceptibility of ciprofloxacin against Salmonella Typhi). Diffusion tests used to determine the susceptibility of organisms isolated from clinical specimens have their own limitations, when equivocal results are obtained, or in prolonged serious infections eg. bacterial endocarditis. In these cases, the quantitation of antibiotic against pathogen needs to be more precise So when in doubt about the sensitivity of pathogen the way to a precise assessment is to determine the minimum inhibitory concentration (MIC) of the antibiotic to the organisms concerned.

II Reagents and lab wares 0.5 McFarland standard, sterile Mueller Hinton agar (pH 7.27.4), sterile Mueller Hinton broth, antibiotic powder, sterile test tubes, pipettes, screw capped flat bottomed bottles (25 ml capacity) and Petri dishes (90 mm diameter). These also include sterile saline (0.85 %) and stock solution of antibiotic. Preparation of stock solutions of antibiotics: The required dilutions of the antibiotics are made as per the table 33-1. Prepare a stock solution containing 2000 µg / ml of the antibiotic to be tested. For example weigh 200 mg of the antibiotic powder and dissolve in 5 ml of distilled water / appropriate solvent. Mix 0.5 ml of this solution with 9.5 ml distilled water (working solution contains antibiotics at a strength of 200 µg / ml-solution A) III Specimens Preparation of suspension of bacteria: Approximately, 4-5 wellisolated colonies of the bacterial strain to be tested are transferred to Tryptic soy broth or BHI broth. The turbidity of the suspension is adjusted to match 0.5 McFarland standards (106 organisms/ml).

PROCEDURE PRINCIPLE In agar dilution method, serial dilutions of the antibiotics are prepared in agar and poured into petri dishes. The dilutions are made in a small volume of water and added to agar which has been melted and cooled to not more than 60°C. Agar dilutions methods have the advantage that it is possible to test a number of organisms on each plate containing an antibiotics solution.

REQUIREMENTS I Equipments Water bath.

Preparation of the agar plate with different concentration of the antibiotics 1 Dispense 2 ml of the diluted antibiotic solution into each of the marked sterile screw capped bottle. Note: It is advisable to start wilh the highest dilution so that single pipette can be used to dispense all the dilutions prepared. 2 Sterile Muller-Hinton agar is cooled and maintained at 50°C – 55°C in a water bath (Table 33-1). 3 Pour this medium (18 ml) into the screw capped bottle containing the different concentration of antibiotic, shake well and pour into sterile petri dish. Note: By this method, exact volume of medium (22.6 ml) is delivered into the screw capped bottles without the danger of the molten agar jellifying during transfer into dilution of the antibiotic (Table 33-1).

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4 Keep the poured plates at 4°C for setting. 5 After the plates have set, dry the plates well in an incubator at 37°C for 30-60 mins. The plates must be dry before performing the test.

Test procedure 1 A grid is marked on the bottom of the plates containing antibiotics. Note: 20 – 25 strains can be tested in plate including the control. 2 A loopful of inoculating loop is calibrated to deliver 0.001 – 0.002 ml (1-2 µl) of the culture. 3 Inoculate the culture on the surface of the medium, indicated by the square marked below. In each case 104 bacteria is delivered to a spot 5 – 8 mm in diameter. Note: Inoculation is done starting with the plates containing highest dilution of the antibiotic. 4 Inoculate a control plate without antibiotics simultaneously as control. 5 Allow the drops to dry and incubate the plates without inverting at 37°C for 16 –18 hours.

QUALITY CONTROL Staphylococcus aureus ATCC 25923, Escherichia coli ATCC 25922, Enterococcus faecalis ATCC 29212, and Pseudomonas aeruginosa ATCC 27853 are used as control strains.

OBSERVATIONS Read the plates for presence or absence of growth. Check the control plate for growth. Control plate must show confluent or near confluent growth. Read the test plate. The concentration at which growth is completely inhibited is considered as the minimum inhibitory concentration (MIC). The organisms are reported sensitive, intermediate or resistant by comparing the test MIC values with that given in the NCCLS table.

RESULTS AND INTERPRETATION The highest dilution of antibiotic showing more than 99% inhibition of growth of bacteria is considered as the minimum inhibitory concentration (MIC) of the bacteria.

Table 33.1 System for preparing dilutions for agar dilution method Antibiotic Solution

+ Sterile Water (Vol)

= Intermediate conc (µg / ml) in tubes

Final conc at 1:10 in agar plates (µg / ml)

Volume

µg/ml

6.4 2 1 1

2000 - A (Stock) 1280 - B 1280 - B 1280 - B

3.6

1280 - B

2 3 7

640 - C 64 320 - D32 160 - E 16

2 1 1

160 - E 160 - E 160 - E

2 3 7

80 - F 8 40 - G 4 20 - H 2

2 1 1

20 - H 20 - H 20 - H

2 3 7

10 - I 1 5 - J 0.5 2.5 - K 0.25

128

KEY FACTS 1 Agar dilution method is a quantitative method for determining the minimum inhibitory concentration (MIC) of the antibiotic against bacteria to be tested. 2 The method is carried out on Muller Hinton agar. 3 It is mainly useful in testing isolates from serious infections like bacterial endocarditis or to verify equivocal results (e.g. intermediate susceptibility of ciprofloxacin against Salmonella Typhi). 4 Selective media should not be used for performing agar dilution method. 5 Electrolyte deficient media also should not be used because it will give false results due to variations in the salt content on action of many antibiotics.

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VIVA 1 What is the break point of an antimicrobial agent? Ans: Break point of an antimicrobial agent is defined as the concentration of that agent that can be achieved in the serum with optimal therapy. 2 What is the automated method used to inoculate the organisms? Ans: Steer’s Replicator is used as the automated method to inoculate the organisms. 3 Mention the merits and demerits of broth dilution method. Ans: Merit of the test is that it is possible to test many organisms on each plate. Demerit is that it is a cumbersome procedure.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. 1997. pp. 1395. Lippincott Williams and Wilkins. 4 Lalitha M.K. Manual on Antimicrobial Susceptibility Testing. Christian Medical College, Vellore, 2004, pp 43. 5 WHO. Guidelines on Standard Operating Procedures for Microbiology. Blood safety and Clinical Technology. Chapter 7: Antimicrobial Susceptibility Testing.

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34

Broth Dilution Method

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Determine antibacterial sensitivity of bacterial isolates by broth dilution method.

INTRODUCTION Broth dilution is also known as tube dilution method. It is another quantitative method for determining the minimum inhibitory concentration (MIC) of the antibiotic against a bacteria to be tested. In this method, serial dilutions of the antibiotics are taken in test tubes and a standardise suspension of the bacterium is inoculated. After incubating overnight , the MIC of the antibiotics is determined by observing the lowest concentration of antibiotics that inhibits growth of the bacteria. The minimum bactericidal concentration (MBC) can also be estimated by this method by subculturing from the lowest concentration of drug that kills the bacteria.

PRINCIPLE A stock solution of antimicrobial agent to be tested is prepared. Two fold dilutions of this solution is prepared in suitable broth. A standard suspension of the organism is inoculated into the medium with one antimicrobial agent -free medium as control. The inoculated media are inoculated at 35-37°C for 18-24 hr. and examined for growth. MIC is taken as the lowest concentration of antimicrobial agent which completely inhibits the growth.

REQUIREMENTS I Equipments Water bath.

II Reagents and lab wares 0.5 McFarland standard, sterile Mueller Hinton broth, antibiotic powder, sterile test tubes. sterile pipettes of 10ml, 5 ml, 2 ml and 1 ml, sterile capped tubes and test tube rack. These also include stock solution of antibiotic. Preparation of stock solutions of antibiotics: The required dilutions of the antibiotics are made as per the table 34-1. Prepare a stock solution containing 2000 µg / ml of the antibiotic to be tested. For example weigh 200 mg of the antibiotic powder and dissolve in 5 ml of distilled water / appropriate solvent. Mix 0.5 ml of this solution with 9.5 ml distilled water (stock solution contains antibiotics at a strength of 200 µg / ml-solution A) III Specimens Preparation of suspension of bacteria: Approximately, 4-5 well isolated colonies of the bacterial strain to be tested are transferred to Tryptic are soy broth or BHI broth. The turbidity of the suspension is adjusted to match 0.5 McFarland standards.

PROCEDURE 1 Serial dilutions of the antimicrobial agent are made in broth and are kept in test tubes. 2 The last tube is kept free of antibiotic and serves as a growth control. 3 Arrange the test tubes in a rack. 4 Standardised suspension of the microorganisms to be tested is inoculated into the tubes. 5 Tubes are incubated at 35-37°C for 18 hours.

QUALITY CONTROL Staphylococcus aureus ATCC 25923, Escherichia coli ATCC 25922, Enterococcus faecalis ATCC 29212, and Pseudomonas aeruginosa ATCC 27853 are used as control strains.

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OBSERVATIONS

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The tubes not showing visible growth are subcultured on solid medium and incubated at 37°C overnight.

At the end of the incubation period the tubes are examined for turbidity. Cloudiness indicates that bacterial growth has not been inhibited by the concentration of antibiotic present in the medium.

Table 34-1 Preparation of stock dilutions of the antibiotic stock solutions

RESULTS AND INTERPRETATION

Stock dilutions of the antibiotic stock solutions can be prepared using the formula:

Minimum inhibitory concentration (MIC) is defined as the highest dilution which inhibits growth judged by lack of turbidity in the tube. The main advantage of the broth dilution method for MIC determination is that it can readily be converted to determine the minimum bactericidal concentration (MBC) also. The highest dilution showing at least 99% inhibition is taken as MBC.

1000 x V X C =W P Where: P =Potency given by manufacturer V= Volume (ml) require C= Final concentration of solution (per ml) W= Weight of antimicrobial to be dissolved in volume V

KEY FACTS 1 Minimum inhibitory concentration (MIC) is defined as the highest dilution which inhibits growth of the bacteria. It is noted by lack of turbidity in the tube. 2 Because very faint turbidity may be given by the inoculum itself, the inoculated tube kept in the refrigerator overnight may be used as the standard for the determination of complete inhibition, 3 Standard strain of known MIC should be tested as control to check the reagents and conditions.

VIVA 1. What is a bacteriostatic agent? Give examples. Ans: Certain antimicrobial agents inhibit the growth by preventing the multiplication of organisms. They do not cause death. These are called bacteriostatic agents. e.g. tetracycline, erythromycin, sulphonamides, etc. 2. What is a bactericidal agent? Give examples. Ans: The drugs that cause irreversible damage to bacteria resulting in death are called bactericidal drugs. e.g. aminoglycosides, penicillin and quinolones. 3. What is minimum inhibitory concentration (MIC)? 4. What is minimum bactericidal concentration (MBC)? Ans: The lowest concentration of the antimicrobial agent that allows < 0.1 % of the original inoculum to survive is called the minimum bactericidal concentration (MBC). 5. Mention merits and demerits of broth dilution agar method. Ans: Merits 1 Simple procedure for testing a small number of isolates, even single isolates. 2 Same tubes can be used for MBC tests also. Demerits There may be non specific turbidity due to the inoculum itself.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. 1997. pp. 1395. Lippincott Williams and Wilkins. 4 Lalitha M.K. Manual on Antimicrobial Susceptibility Testing. Christian Medical College, Vellore, 2004, pp 43. 5 WHO. Guidelines on Standard Operating Procedures for Microbiology. Blood safety and Clinical Technology. Chapter 7: Antimicrobial Susceptibility Testing.

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35

Epsilometer Test (E-test)

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to:

I Reagents and lab wares

1 Determine antibacterial sensitivity of bacterial isolates by Epsilometer test (E-test), an automated system for measuring the minimum inhibitory concentration (MIC) of the bacteria.

Commercially available E-test strips, 0.5 McFarland standards, sterile Mueller Hinton agar plates (150 or 90 mm with a depth of 4 mm). In a 90 mm plate, a single antibiotic strip can be tested. In a 150 mm plate, at least 4 antibiotic strips can be tested. These also include foreceps, 0.85% saline for inoculum preparation, and sterile swabs

INTRODUCTION The E-test is an automated system for measuring the MIC of the bacterial isolate. It is a very simple test to perform MIC of the bacterial isolate as compared to the other techniques like broth and agar dilution methods which are technically cumbersome. An elliptical zone of growth inhibition is seen around the strip after incubation. The MIC is read from the scale at the intersection of the zone with the strip. It is easy to interpret the result of MIC.

II Specimens Preparation of suspension of bacteria: Inoculate peptone water with test organism and incubate at 37 °C for 3-4 hours. The turbidity of the suspension is adjusted to match 0.5 McFarland standards.

PROCEDURE Opening an E-test package

PRINCIPLE The E-test is based on the principle of disc diffusion where the antibiotic diffuses into the medium when the strip is placed on the medium. The E-test is a plastic strip (5 x 50 mm; antibiotic carrier) with a continuous gradient of antibiotic immobilized on one side and MIC interpretative scale corresponding to 15 two fold MIC dilutions on the other side. A predefined antibiotic gradient is immobilized on the surface opposite the MIC scale. When transferred to the agar, the continuous antibiotic gradient established under the strip remains stable over a period covering the critical times of most microorganisms subjected to susceptibility testing. This method produces a means for producing MIC data in those situations in which the level of resistance can be clinically important. e.g. penicillin or cephalosporins against Streptococcus pneumoniae.

1 Remove the package stored at – 20°C or - 70°C. 2 Equilibrate at room temperature. This takes approximately 30 minutes if stored at – 20° C and approximately one hour if stored at – 70° C. Ensure all the moisture has evaporated before opening. 3 Inspect the package for holes or cracks. Do not use if damaged. 4 Cut along the broken line at the top of a blister. Do not cut in between the blisters. 5 Tip the strips out of the opening slightly and take them out with forceps. 6 If strips stick together, twist them apart with your fingers. 7 Touch only the handle, i.e., the area labeled E. 8 Place the strips to be used into a dry clean petri dish.

Application of strips 1 Apply E-test strips with forceps. Ensure the MIC scale is facing upwards i.e. towards the opening of the plate.

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2 Ensure that the agar surface is dry before swabbing it. Dip a swab in the inoculum, remove excess fluid and swab the entire agar surface evenly in 3 directions. 3 Allow the agar surface to dry for 10 minutes to 15 minutes on the bench or in the incubator. 4 Open the E-test package and place the strips in a dry petri dish. 5 Apply the strips to the agar surface with a forceps. Always apply the strip with the MIC scale facing the opening of the plate. Do not apply it upside down. Note: Be firm when applying the strip. Once applied, do not move the strip. 6 Use templates to position 4 to 6 strips on a 150 mm plate or one to two strips on a 90 mm plate. 7 Place the handle of the strip closest to the rim of the plate. Note: Always store unused strips in airtight containers at 20°C or -70°C. 8 Incubate at 37°C for 18-24hours.

QUALITY CONTROL Package labels for each antibiotic will carry performance and reproducibility data. NCCLS QC ranges and interpretive guidelines.

OBSERVATIONS After incubation an elliptical zone of growth inhibition is seen around the strip. The MIC is read from the scale at the intersection of the zone with the strip (Fig. 35-1).

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FIGURE 35-1 E-Test.

RESULTS AND INTERPRETATION Read plates after the recommended incubation period only if sufficient growth is seen and the inhibition eclipse is clearly visible. Read the MIC where the ellipse intersects the scale. Always read the end point at complete inhibition of all growth including hazes and isolated colonies. Since E- test comprises a continuous gradient, MIC values in between two – fold dilutions can be obtained. Always round up these values to the next two-fold dilution before interpretation. For example: If ampicillin breakpoints are given as S=1, I = 2, R=4 µg/ml, then an E-test MIC of 1.5 µg/ml is rounded up to 2 µg/ml and the category reported as Intermediate (I)

KEY FACTS 1 The E-test strips have to be placed in proper orientation. Placing strips upside down on the agar will alter the results. 2 Diffusion of antibiotics begins immediately after placement of the strips, which cannot be moved once it has touched the agar. 3 Read plates after the recommended incubation period only if sufficient growth is seen and the inhibition eclipse is clearly visible. 4 Read the MIC where the ellipse intersects the scale. 5 Always store unused strips in airtight containers at -20°C or -70°C.

VIVA 1 What is the principle of the E-test? 2 What is the advantage of E-test? 3 How is the result for MIC interpreted?

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. 1997. pp. 1395. Lippincott Williams and Wilkins. 4 Lalitha M.K. Manual on Antimicrobial Susceptibility Testing. Christian Medical College, Vellore, 2004, pp 43. 5 WHO. Guidelines on Standard Operating Procedures for Microbiology. Blood safety and Clinical Technology. Chapter 7: Antimicrobial Susceptibility Testing.

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UNIT

VI Immunology

Lesson Lesson Lesson Lesson Lesson Lesson Lesson Lesson Lesson Lesson Lesson Lesson Lesson Lesson

36 37 38 39 40 41 42 43 44 45 46 47 48 49

Introduction Bacterial Agglutination Test Blood Grouping Latex Agglutination Test Co-agglutination Test Widal Test Weil Felix Test Anti-Streptolysin O (ASLO) Test VDRL Test Radial Immunodiffusion Test Immunoelectrophoresis Test Counter-current Immunoelectrophoresis Test Indirect Haemagglutination Test Immunofluorescence Test Enzyme-Linked Immunosorbent Assay

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Introduction Serological tests are widely used for diagnosis of many infectious diseases including bacterial, viral, fungal and parasitic. These tests may be agglutination, precipitation, neutralization, etc. with a variation in their sensitivity and specificity. List of few common serological tests are mentioned in the table.

Table Applications of various tests used in a microbiology laboratory TYPE OF THE TEST

NAME OF THE TEST

APPLICATION

Slide agglutination tests.

Bacterial agglutination test.

For confirmation of identification of the bacterial isolates by using specific antisera. Shigella agglutination. Salmonella agglutination. Vibrio agglutination. Streptococcus Lance field’s grouping. For blood group antigen detection A, B, AB and O. For antibody detection against Salmonella Typhi S. Paratyphi A S. Paratyphi B S. Paratyphi C.

Blood grouping. Widal test.

Tube agglutination tests.

Standard agglutination test. Cold agglutination test. Indirect haemagglutination test.

Passive agglutination tests.

Latex agglutination test.

Co-agglutination test.

For antibody detection against Brucella species. For antibody detection against Mycoplasma pneumoniae. For antibody detection in: Amoebiasis Lymphatic filariasis Echinococcosis Toxoplasmosis Rickettsial infection. For antigen detection in infections caused by: Streptococcus pneumoniae Haemophilus influenzae Cryptococcus neoformans Echinococcus granulosus. For antigen detection in: Cryptococcosis Echinococcosis Lymphatic filariasis

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Heterophile agglutination tests.

Weil-Felix test. Paul-Bunnel test.

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For heterophile antibody detection in Scrub typhus Endemic typhus. Epidemic typhus. For heterophile antibody detection in Infectious mononucleosis (EBV).

Neutralization test

ASLO test.

For detection of antibodies (Anti-streptolysin) in acute rheumatic fever.

Precipitation tests

Ring test.

For antigen detection in infections caused by: Brucella (Milk ring test) B.anthracis (Ascoli’s test) Toxoid precipitation for diphtheria. Slide flocculation test for demonstration of reaginic antibodies in syphilis. For detection of: a-Fetoprotein N. meningitidis antigen HBs antigen.

Tube test. VDRL test. Gel diffusion test.

Immunofluorescence test

Direct immunofluorescence test.

For antigen detection in infections caused by: Respiratory syncytial virus Measles Mumps Rabies Influenza Indirect immunofluorescence test. For antibody detection in: Toxoplasmosis Amoebiasis.

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36

Bacterial Agglutination Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the application of slide agglutination reaction for the identification of bacteria.

only small quantities of culture are available. The slide agglutination tests have many uses. They are used for confirmatory identification of Salmonella, Shigella and Vibrio isolates, identification of Bordetella pertussis and typing of streptococci (e.g. Streptococcus Lance field’s grouping) and pneumococci. The slide agglutination test is rapid and convenient.

INTRODUCTION PRINCIPLE Agglutination is an example of antigen-antibody reaction in which antigen is particulate or insoluble in nature. The soluble antigens can be tested in agglutination reaction by coating them with carrier particles such as red blood cells, bacteria or inorganic particles such as the latex. When a particulate antigen is mixed with its antibody in the presence of electrolytes at a suitable temperature and pH, the particles are clumped or agglutinated. Antibodies cause agglutination by binding to antigens on the particles; they help to neutralize the slight negative charge that particles in solution normally carry (the zeta potential). This allows the particles to approach each other. IgM immunoglobulins actually function as a bridge between two particles, with one of the five subunits binding to one particle, and another subunit binding to another particle. As in the case with precipitation reaction, more quantity of antibody can actually inhibit agglutination. In this case, each particle will be completely covered with antibodies, and they will not clump together. This phenomenon is known as prozone reaction. Not all antibodies can agglutinate particles. Antibodies, which cannot cause agglutination, are called “incomplete” antibodies. This is probably because of restricted movement in the hinge region of the immunoglobulin. IgG4 antibodies are an example of incomplete antibodies. The slide agglutination is a frequently used procedure for the identification of many bacterial isolates from clinical specimens. This method is also used for blood grouping and cross matching. The test is performed on a glass slide, hence called slide agglutination test. This method is useful where

In the slide agglutination test a drop of the appropriate specific antiserum is added to a smooth, uniform suspension of a bacterial isolate from the clinical specimens resulting in agglutination. A positive reaction is identified by the clumping together of bacteria and clearing of the drop. The reaction is facilitated by mixing of the bacterial colony and the antiserum with a loop. The agglutination reaction occurs instantly or within seconds.

REQUIREMENTS I Reagents and glass wares Glass slides, bacteriological loop, glass marking pencil, saline (0.85%), specific antiserum against the bacterium(e.g., antiserum against Salmonella Typhi, Shigella flexneri or Vibrio cholerae) to be tested. II Specimen Pure 24 hour growth of bacteria (e.g., Salmonella Typhi, Shigella flexneri or Vibrio cholerae) from solid media preferably from non-blood agar plates (Examples : nutrient agar, Muller-Hinton agar).

PROCEDURE 1 Take a clean glass slide.

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2 Mark it into two halves by a glass marking pencil and label them as test and control. 3 Place a drop of saline in both the halves. 4 Pick up the colonies of Salmonella to be tested from agar culture and gently emulsify with drops of saline in both the halves by loop. 5 Add a drop of specific antisera to the bacterial suspension in the half labeled as test and mix. 6 Place another drop of saline in the half of the slide labeled as control, and mix. 7 Gently rock the slide, back and froth for 2 minutes. 8 Observe the clumping of the bacterial suspension in the test.

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the test reagent as compared to control reagent is indicative of a positive reaction. Absence of reaction with a test reagent indicates a negative reaction irrespective of any reaction with control organism.

RESULTS AND INTERPRETATION Positive agglutination: Clumping in the test half and no clumping in the control half. It identifies specific bacteria (Fig. 36-1). Negative agglutination: No clumping either in the test half or control half. It shows either antiserum is not good or bacteria and the antisera are not specific to each other. Auto agglutination: Clumping in both test and control halves. Test is fallacious, hence discard the result.

On the same slide a control consisting of the bacterial suspension in saline without the antisera is used to ensure that bacteria is not auto agglutinable.

OBSERVATIONS Observe the test mixture for agglutination by naked eye observation, but sometimes it may require confirmation under the microscope. A markedly stronger agglutination reaction in

POSITIVE

NEGATIVE

FIGURE 36-1 Slide agglutination test.

KEY FACTS 1 Fresh young cultures are always used for the test. 2 Check for the auto agglutination of the test organism. 3 On the same slide a control consisting of the bacterial suspension in saline without the antisera is used to ensure that bacteria are not auto agglutinable.

VIVA 1 Discuss application of slide bacterial agglutination tests. 2 How do you check for the auto agglutination of the test organism?

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

37

Blood Grouping

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to: 1 Demonstrate blood grouping by agglutination reaction.

1 Take three different slides and label the slides as A, B, and D. 2 Clean middle finger of the left hand with the spirit and allow it to dry. 3 Prick the finger with sterile lancet. 4 Collect 3 drops of blood on three different slides labeled as A, B, and D. 5 Add a drop of antiserum A to A and anti B to B anti D to D. 6 Mix with an applicator stick. Mix the samples on the slide by gentle rocking for about two minutes. 7 Examine each zone for agglutination of RBCs. 8 Record result of the test immediately before the drop dries out.

INTRODUCTION The ABO system contains four blood groups and is determined by the presence or absence of two distinct antigens, A and B, on the surface of erythrocytes. Red cells of group A carry antigen A, cells of group B antigen B and cells of group AB have both A and B antigens, while group O cells have neither A nor B antigen. The four groups are also distinguished by the presence or absence of two distinct isoantibodies in the serum. The serum contains the isoantibodies specific for the antigen that is absent on the red cell. Blood group antigens are inherited according to Mendelian laws. Their synthesis is determined by allelomorphic genes A, B and O. Genes A and B give rise to the corresponding antigens, but O is an amorph and does not produce any antigen. Group O is the commonest group and group AB is the rarest. In India the distribution is O – 40%, A – 22%, B – 33%, and AB – 5%. O group population is called universal donors and AB is called universal recipients.

PRINCIPLE When a drop of anti A / anti B or anti Rh antibody is added to a drop of blood, the antibody binds with its specific antigen present on the RBCs and causes agglutination of the RBCs.

REQUIREMENTS I Reagents and lab wares Blood group antisera (Anti-A, anti-B, anti-D or Rh antiserum), glazed white ceramic slide, applicator stick and lancet. II Specimen Blood with anticoagulant or finger prick blood.

QUALITY CONTROL Blood group antisera (Anti-A, anti-B, anti-D or Rh antiserum) must be checked with known blood before to test.

OBSERVATIONS Observe the test mixture for clumping. Check for autoagglutination of test RBCs.

RESULTS AND INTERPRETATION A clear agglutination indicates positive result. If the test sample is agglutinated with anti A antibody then the blood is Group A. If the test sample is agglutinated with anti B antibody then the blood is Group B. If the test sample is agglutinated with anti A and anti B antibodies both, then the blood is Group AB. If the test sample is not agglutinated with anti A and anti B antibodies, then the blood is Group O.

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KEY FACTS 1 Autoagglutination of RBCs during the test procedure should be checked. 2 Record result of the test immediately before the drop dries out.

VIVA 1 What exactly does it tell you about a person’s blood cells if you know that they are of blood type A, B, AB, or O? Ans: It shows the presence or absence of two distinct antigens, A and B, on the surface of erythrocytes. 2 Explain what blood types a person of type A, B, AB, or O can receive and why? Ans :Red cells of group A carry antigen A, cells of group B carry antigen B and cells of group AB have both A and B antigens, while group O cells have neither A nor B antigen. The serum contains the isoantibodies specific for the antigen that is absent on the red cell. Immune isoantibodies may develop following ABO incompatible transfusion. Red cells also carry another antigen that reacts with rabbit serum to Rhesus monkey erythrocytes called Rhesus or Rh factor. Rh-negative population contains anti Rh antibodies in their blood. Hence a person should not receive blood, which contains isoantibodies to own RBC. O group population is called universal donors since their blood does not contain any blood group antigens (A, B and AB). AB group is called universal recipients due to absence of isoantibodies.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

38

Latex Agglutination Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the application of latex agglutination test for detection of soluble antigen.

INTRODUCTION Latex agglutination test is an example of slide agglutination test in which polystyrene spheres such as latex particles have been used as carrier particles for coating with antigens. In this test both soluble proteins and non-protein antigens can be bound on the surface of latex beads, which then can be used to detect antibodies by demonstrating the agglutination of the antigen-carrying latex particles. The latex agglutination test is used for demonstration of antibodies on a variety of bacterial, parasitic and fungal infections. The LAT are widely used for diagnosis of typhoid fever, syphilis, infectious mononucleosis, amoebiasis and hydatid diseases. The LAT can also be used for detection of soluble antigen by using latex particles coated with the specific antibodies. The LAT is used to demonstrate rheumatoid factor (RF) in the serum, Streptococcus pneumoniae and Haemophilus influenzae and cryptococcal antigen in the cerebrospinal fluid for diagnosis of meningitis, Clostridium difficile antigens in the stool for diarrhoea and hydatid antigen in the serum for diagnosis of hydatid disease. In this exercise you will perform the LAT for detection of antigen in the serum for diagnosis of hydatid disease.

PRINCIPLE In the LAT for detection of hydatid antigen in the serum, the latex particles are coated with specific polyclonal hydatid antibodies. Then hydatid antibody coated latex particles are mixed with serum to be tested for the antigen, If serum contain antigen, then the latter combines with the antibodies on surface

of the latex particles and form visible agglutination or clumping of the particles.

REQUIREMENTS I Reagents and lab wares Hydatid antibody-coated latex bead, glass slides, and applicator stick. II Specimen Serum sample to be tested.

PROCEDURE 1 Take a clean glass slide. 2 Mark it into two halves by a glass marking pencil and label them as test and control. 3 Add a drop of test serum in the test half, and a drop of saline in the control half. 4 Add a drop of hydatid antibody-coated latex reagent to the serum in the half labeled as test and mix. 5 Place another drop of hydatid antibody-coated latex reagent in half of the slide labeled as control, and mix. 6 Gently rock the slide, back and froth for 2 minutes. 7 Observe the clumping of the latex reagent in the test.

QUALITY CONTROL The LAT is performed with a known positive and negative hydatid sera, every time the test is performed with the test sera.

OBSERVATIONS Observe the test half for agglutination of latex reagent. Observe the control half for autoagglutination.

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RESULTS AND INTERPRETATION Agglutination of latex reagents with the sera, and absence of any agglutination in the control half indicates the LAT to be positive and shows the presence of hydatid antigen in the serum (Fig. 38-1). Absence of agglutination either in the test half or in the control half indicates the test to be negative and shows the absence of hydatid antigen in the serum.

NEGATIVE

POSITIVE

FIGURE 38-1 Latex agglutination test.

KEY FACTS 1 Latex agglutination test is an example of slide agglutination test. 2 The latex agglutination test is used for demonstration of antibodies on a variety of bacterial, parasitic and fungal infections. 3 The LAT can also be used for detection of soluble antigen by latex particles coated with the specific antibodies.

VIVA 1 List application of the LAT for diagnosis of infectious diseases by demonstration of antibodies in the serum. 2 List application of the LAT for diagnosis of infectious diseases by demonstration of antigens in the serum.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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39

Co-Agglutination Test

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to: 1 Demonstrate the application of Co-agglutination test for detection of soluble antigen.

1 Take a clean glass slide. 2 Mark it into two halves by a glass marking pencil and label them as test and control. 3 Add a drop of test CSF in the test half, and a drop of saline in the control half. Note: The CSF specimen has to be absorbed with stabilized SAPA cells (not coated with specific pneumococcal antibody) to prevent nonspecific agglutination with human IgG. 4 Add a drop of pneumococcal antibody-coated Co-A reagent to the serum in the half labeled as test and mix. 5 Place another drop of pneumococcal antibody-coated Co-A reagent in the other half of the slide labeled as control, and mix. 6 Gently rock the slide, back and froth for 2 minutes. 7 Observe the agglutination of the bacteria in the test.

INTRODUCTION The Co-agglutination (Co-A) test is a simple slide agglutination test. The test is being used for detection of specific antigen in the serum, cerebrospinal fluid (CSF), urine and other body fluids. Commercial Co-A kits are now available for detection of haemophilus, meningococcal and pneumococcal antigens in the CSF, and salmonella antigen in the serum. The kits are also available for identification of Neisseria gonorrhoeae and sero grouping of Staphylococcus aureus. In this exercise you will perform the Co-A for detection of pneumococcal antigen in the CSF for diagnosis of meningitis.

PRINCIPLE Co- agglutination test means cocci- mediated agglutination. In this test certain strains of S. aureus organisms (Cowan’s I strain) containing a large amount of an antibody binding Protein A (SAPA) in their cell walls is utilized as a carrier particle. These cocci on mixing with the specific antibodies (raised against Streptococcus pneumoniae to be detected in the CSF) bind IgG non-specifically through the Fc region leaving specific Fab sites free. The subsequent reaction of Fab with pneumococcal antigen in the test CSF is visualized by clumping.

REQUIREMENTS I Reagents and lab wares Co-agglutination reagent, glass slides, and applicator stick. II Specimen CSF sample to be tested.

QUALITY CONTROL The Co-A is performed with a known pneumococcal antigen positive and negative CSF, every time the test is performed with the test sera.

OBSERVATIONS Observe the test half for agglutination of Co-A reagent. Observe the control half for auto agglutination.

RESULTS AND INTERPRETATION Agglutination of the Co-A reagents with the CSF , and absence of any agglutination in the control half indicates the Co-A to be positive and shows the presence of pneumococcal antigen in the CSF. Absence of agglutination either in the test half or in the control half indicates the test to be negative and shows the absence of pneumococcal antigen in the CSF.

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KEY FACTS 1 The Co-agglutination test is an example of slide agglutination test. 2 S. aureus organisms (Cowan’s I strain) containing a large amount of an antibody binding Protein A (SAPA) in their cell walls is utilized as a carrier particle. 3 The Co-A test is used for demonstration of antigens in the serum, urine and CSF in a variety of infections. 4 The commercial Co-A reagents are available for identification of N. gonorrhoeae and serogrouping of S. pyogenes A, B, C, D or G. 5 The reagents are also available commercially for identification of meningococcal, pneumococcal and haemophilus antigen in the CSF for diagnosis of meningitis.

VIVA 1 Explain the principle of the Co-agglutination test. 2 What is the strain used in Co-agglutination test? 3 List the uses of Co-agglutination test.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

40

Widal Test

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to:

I Equipments Water bath (Fig. 40-1).

1 Perform Widal test for diagnosis of enteric fever.

INTRODUCTION Widal test is an example of tube agglutination test used widely for diagnosis of enteric fever. Enteric fever is caused by Salmonella Typhi, and by Salmonella Paratyphi A, B and C. In India enteric fever is caused commonly by S. Typhi, and by S. Paratyphi A, rarely by S. Paratyphi B. In Widal test serum of the patients is tested for antibodies against H and O antigens of S. Typhi, and S. Paratyphi A and B.

II Reagents and glass wares Small test tubes, test tube racks, and pipettes. Saline (0.85 % NaCl), and salmonella antigen suspensions. These antigens can be prepared in the laboratory or commercially prepared suspensions are available from the Central Research Institute, Kasauli, (Himachal Pradesh). The antigens are stored at 4 °C before use. III Specimen Serum sample to be tested Note: Serum samples may be stored at 4 °C for 10-15 days before testing. They may be stored at –20 °C for the period beyond 2 weeks.

PRINCIPLE Widal test detects antibodies in serum against H and O suspensions of S. Typhi and against H suspension of S. Paratyphi A and S. Paratyphi B. Since S. Paratyphi C infections are not found in India, serum antibodies against this species are not tested. Since “O” antigens are related to each other, the O antigen of S. Typhi is only tested, while H antigens of S. Typhi, S. Paratyphi A and S. Paratyphi B are tested separately. In this test the serum to be tested are diluted in series in test tubes and antigen suspensions of salmonella are added to it. Four rows of such dilutions are prepared and to that antigen suspensions of O antigen of S. Typhi and H antigens of S. Typhi, S. Paratyphi A and S. Paratyphi B are added. The test is then incubated in water bath at 37°C and is checked for agglutination by examining the bottom of the tubes for sediment. The titre of patient serum is calculated for each salmonella suspension as the highest dilution producing visible agglutination.

PROCEDURE Preparation of master dilution of the serum 1 Add 2.3 ml of saline in a test tube. 2 Add 0.2 ml of serum to the test tube, which makes the serum dilution as 1: 12.5. 3 Patient’s serum is tested in a series of dilutions against each of the different antigen suspensions.

Performance of the test 1 Arrange 4 rows of test tubes, each row containing 7 test tubes. 2 Label the row 1 as TO, 2 as TH, 3 as AH and 4 as BH. 3 Add 0.2 ml of diluted serum from master dilution into the first and second tubes in all rows of the tubes, TO, TH, AH and BH. 4 Add 0.2 ml of saline to all the tubes from 2nd to 7th in all the rows including the 7th control tube.

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5 In the row labeled as TO, mix and transfer 0.2 ml of serum from 2nd tube to the 3rd, then from 3rd to 4th, and so on through the 6th tube. Discard 0.2 ml from the 6th tube. 6 Follow the same dilutions in the rows labeled as TH, AH and BH. 7 Add 0.2 ml of TO antigens to tubes from 1st to 7th in the row TO. 8 Add 0.2 ml of TH antigens to tubes from 1st to 7th in the row TH. 9 Add 0.2 ml of AH antigens to tubes from 1st to 7th in the row AH. 10 Add 0.2 ml of BH antigens to tubes from 1st to 7th in the row BH. Note: The final dilution of the sera after addition of antigen are 1:25, 1;50, 1;100, 1:200, 1:400, 1:800. 11 Incubate the racks in water bath at 37 °C for 18 hrs.

QUALITY CONTROL 7th tube in each row acts as a negative antigen control. The same dilutions of a known positive serum is tested with TO, TH, AH and BH antigens.

The last tube showing agglutination is considered as the end point. The reciprocal of the dilution is considered as the titre (e.g., If the dilution of the last tube showing agglutination is 1: 200, then the titre is 200).

RESULTS AND INTERPRETATION 1 A progressive rise in the titer between first and third week after onset of fever is highly significant. 2 A positive or a negative result in a single test is not significant. 3 Since the antibodies are detected only after 7 days to 10 days of illness, test should be done later. 4 The serum of some uninfected subjects causes agglutinations at dilutions of about 1:50, so titers are considered significant when agglutination occurs in serum dilution above 100. 5 H agglutinins tend to persist longer than O agglutinins. 6 O antibody titre rise indicates recent infection. 7 Persons immunized with TAB vaccine may show high titers of antibodies to all the antigens and so only a marked rise in titer is considered significant. 8 Early treatment with antibiotics will alter antibody response.

OBSERVATIONS Observe the test tubes after 18 hours of incubation in waterbath at 56°C for agglutination and note the result. The results are read by viewing the tubes under good light against a dark background with the aid of a magnifying lens. In case of H agglutinins large loose, cotton and wooly clumps are formed and with O agglutinins only small granules are formed at the bottom of the tube. If necessary, the tubes can be gently rotated, to swirl up granules from the deposit. The titer of the serum is the highest dilution of serum giving visible agglutination. No agglutination is seen as a small compact deposit (button) formation.

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FIGURE 40-1 Water bath.

VIVA 1. What are the advantages and disadvantages of Widal test? Ans: Advantages: Simple and inexpensive test. Sensitive test for the diagnosis of typhoid in endemic areas. Useful for cases in children, who have a low prevalence of pre-existing antibody. Disadvantages: Less specific. A positive or a negative result in a single test is not significant. The antibodies are detected only after 7 to 10 days of illness. The serum of some uninfected subjects causes agglutinations (False positive reactions). Persons immunized with TAB vaccine may show high titers of antibodies to all the antigens. Cases treated early with chloramphenicol may show a poor agglutinin response.

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KEY FACTS 1 In India enteric fever is caused commonly by S. Typhi, and S. Paratyphi A, rarely by S. Paratyphi B. 2 Since “ O” antigens are related to each other, the O antigen of S. Typhi is only tested, while H antigens of S. Typhi, S. Paratyphi A and S. Paratyphi B are tested separately. 3 The last tube showing agglutination is considered as the end point. 4 The reciprocal of the dilution is considered as the titre(e.g., If the dilution of the last tube showing agglutination is 1: 200, then the titre is 200).

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

41

Weil-Felix Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the cross-reacting antibodies to Rickettsial antigens using Proteus strains OX-19, OX-2 of Proteus vulgaris and OX-K of P. mirabilis by Weil-Felix test.

II Reagents and glass wares Small test tubes, test tube racks, and pipettes. Saline (0.85 % NaCl), and P. vulgaris OX-19, OX-2 and P. mirabilis OX-K antigens. The antigens are stored at 4°C before use. III Specimen Serum sample to be tested.

INTRODUCTION PROCEDURE Weil-Felix is a serological test used for diagnosis of Rickettsial infection by demonstration of antibodies in the serum. In this test, certain strains of Proteus are used instead of specific rickettsial pathogens as antigens. This test was developed from the observation that certain strains of Proteus isolated from the urine of patients with epidemic typhus were agglutinated by the sera of the typhus patients. This test has been used for presumptive serological evidence of Rickettsial disease.

PRINCIPLE It is an agglutination test for cross-reacting antibodies. It is based on the principle that many patients infected with one of the Rickettsia produce antibodies that can agglutinate certain strains of bacteria of genus Proteus because of the presence of a common alkali stable carbohydrate antigen. Rickettsia prowazaki and R. mooseri share alkali stable carbohydrate antigens with P. vulgaris OX 19, R. tsutsugamushi with P. mirabilis OX K and R.rickettsi, and R.conori with P. vulgaris OX 2.

REQUIREMENTS I Equipments Water bath.

Preparation of master dilution of the serum 1 Add 2.7 ml of saline in a test tube. 2 Add 0.3 ml of serum to the test tube which makes the master serum dilution as 1: 10.

Performance of the test 1 Arrange 3 rows of test tubes, each row containing 7 test tubes. 2 Add 1 ml of saline to all the tubes. 3 Add 1 ml of diluted serum from master dilution into the first tubes in all the three rows of the tubes. 4 Make doubling dilutions in the first row from the first tube(serum dilution 1:20) and discard 1 ml from the 6th tube (serum dilution 1:640). Note: Leave the 7th tube as control without serum. 5 In the same way, make doubling dilutions in the second and third row tubes. 6 Add 0.5 ml of concentrated P. vulgaris OX-19 in to all 7 tubes in the first row of tubes. 7 Add 0.5 ml of concentrated P. vulgaris OX-2 in to all 7 tubes in the second row of tubes. 8 Add 0.5 ml drop of concentrated P. mirabilis OX-K in to all 7 tubes in the third row of tubes. 9 Incubate the racks in water bath at 37°C for 2 hr., followed by reincubating at 4°C for 18 hours.

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QUALITY CONTROL 7th tube in each row acts as a negative antigen control. The same dilutions of a known positive serum are tested with Proteus OX 19, OX 2 and OX K antigens.

OBSERVATIONS Observe the test tubes after overnight incubation for agglutination and note the result. The results are read by viewing the tubes under good light against a dark background with the aid of a magnifying lens. Complete agglutination is demonstrated by complete clearing of the supernatant fluid and the formation of white flocculent masses in the bottom of tubes

The last tube showing agglutination is considered as the end point. The reciprocal of the dilution is considered as the titre(e.g., If the dilution of the last tube showing agglutination is 1: 640, then the titre is 640).

RESULTS AND INTERPRETATION A serum titer of 1: 80 and above is considered significant titer but four fold or greater increase of antibody between acute and convalescent sera is considered diagnostic. Agglutination with Proteus OX 19 and OX 2 is suggestive of spotted fever. Agglutination with Proteus OX 2 is suggestive of murine typhus. Agglutination with Proteus OX K is suggestive of scrub typhus.

KEY FACTS 1 Since the Weil-Felix antigens also react with Proteus antibodies, false positive reactions may occur in urinary tract infections with Proteus, in leptospirosis, in Borrelia infections and in severe liver disease. 2 Proper precautions should be taken for standardization of antigens. It should not be standardized against sera from rabbits immunized with homologous strain of Proteus species, but with sera derived from patients infected with rickettsiae. 3. The test is not helpful for the detection of antibodies in rickettsial pox, trench fever or Q fever as these persons do not develop Proteus agglutinins.

VIVA 1 What are heterophile antibodies? Ans: Antibodies produced in certain clinical conditions have the ability to cross react with more than one antigen called heterophile antibodies 2 What is the principle of Weil-Felix reaction? 3 List other tests detecting heterophilic antibodies Ans: a. Streptococcus MG agglutination test for the diagnosis of atypical pneumonia. b. Paul-Bunnel test for the diagnosis of Epstein-Bar virus infections and c. cold agglutination test for the diagnosis of primary atypical pneumonia.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

42

Anti-Streptolysin O (ASLO) Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate neutralizing antibodies to Streptococcus pyogenes haemolysin O antigen.

INTRODUCTION Str. pyogenes produces several exotoxins and enzymes, which contribute to its virulence. Streptococci produce two haemolysins, namely streptolysin O and streptolysin S. Streptolysin O is oxygen labile and heat labile. It is inactive in the oxidized form but may be reactivated by treatment with mild reducing agents. It is lethal on intravenous injection into animals and has a specific cardiotoxic and leucotoxic activity. Streptolysin O is antigenic and elicits the production of specific antibodies against the antigen in an infected human host. This antibody is known as antistreptolysin, it regularly appears in sera following streptococcal infection. Hence, demonstration of antistreptolysin in the serum is an indirect indicator of infection. Many methods are available for detection of antistreptolysin in the serum. Streptolysin S is not antigenic, hence do not elicit production of any antibodies.

PRINCIPLE When streptolysin O in its reduced form is added to red blood cells hemolysis occurs. If a patient’s serum contains antistreptolysin O antibodies, antigen-antibody reaction takes place and no hemolysis takes place.

II Reagents and lab wares Antigen (Streptolysin O), phosphate buffered saline pH 6.36.5, 2.5% RBC’s in Alsever’s solution (human blood group O or rabbit erythrocytes), microtitre plates, pipette, test tubes. III Specimen Serum.

PROCEDURE Serum dilution 1 The initial serom dilutions of 1:10, 1:60 and 1:85 are prepared in test tubes. Subsequent dilutions are carried out in microtitre plates. 2 Both serum and buffer should be placed at room temperature when preparing dilution. 3 Standard serum of know titer is included in each days run to serve as positive control.

Test procedure 1 Label microtiter plate so that each specimen is alligned in two rows (1:60 row and 1:85 row) with six wells per row. 2 Prepare serial two fold dilutions through sixth well. Serum dilutions for each serum sample are first row - 1:60, 1:120, 1:240, 1:480, 1:960, 1:1920; second row - 1:85, 1:170, 1:340, 1:680, 1:1360, 1:2760. 3 Add 0.025ml of antigen to all tubes, gently agitate, and incubate at 37°C for 15 minutes. 4 Add 0.025 ml of cold 2.5% red blood cells to all wells and incubate in water bath at 37° C for one hour.

QUALITY CONTROL A cell control, antigen control and a standard serum control always should be used with the test.

REQUIREMENTS I Equipments Water bath.

OBSERVATIONS Observe the microtitre plates for presence or absence of hemolysis.

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RESULTS AND INTERPRETATION Reading is taken by observing for button formation and hemolysis in microtitre plate.

The titer of ASLO is the highest serum dilution causing no hemolysis. Significant titer is considered to be 200 or more.

KEY FACTS 1 Streptolysin O is antigenic and elicits the production of specific antibodies against the antigen in an infected human host. 2 Streptolysin S is not antigenic; hence do not elicit production of any antibodies.

VIVA 1 Name the hemolysins produced by Streptococci. 2 Explain the principle of the LAT test.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

43

VDRL Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform the VDRL test. 2 Demonstrate the presence of reaginic antibodies in patient’s serum.

test. When cardiolipin antigen combines with reaginic antibodies in patient’s serum, it forms visible floccules. This test is also used to detect antibodies in the CSF. The biological false positive reaction is a noted problem associated with the VDRL test (Box 43-2).The advantages and disadvantages of the VDRL test are listed in the table 43-1.

REQUIREMENTS INTRODUCTION When soluble antigen and specific antibodies in the serum are combined in the right proportions, they will bind to each other to form an insoluble complex which results in visible precipitations. The precipitates settle at the bottom of the container. The precipitation reaction can be demonstrated in the liquid medium as well as in the solid media such as in gels. The process of precipitation in the gel can be hastened by migration of antigens and antibodies under an electric field. Temperature, pH and concentration of sodium chloride in the medium influence the reaction of precipitation. Often the precipitate fails to settle down at the bottom of the container but remains suspended as floccules, such reaction is known as flocculation. Venereal Disease Research Laboratory (VDRL) test is an example of flocculation test to detect reaginic antibodies in serum of patients suffering from syphilis. VDRL –ELISA is a modification of the VDRL test (Box 43-1).

I Equipments Water bath and VDRL shaker. II Reagents and glass wares VDRL antigen (It contains cardiolipin, 0.3%, lecithin, 0.24% and cholesterol, 0.9%), buffered saline solution, saline, glass stoppered bottles, VDRL glass slides (2”X3”) with depressions of 14 mm diameter each, tubes, pipettes and racks. Preparation of buffered saline solution: This is prepared by mixing 0.5 ml of formaldehyde, 3.037 gm of disodium hydrogen phosphate (Na 2HPO 4, 2H 2O), 0.170 gm of potassium dihydrogen phosphate (KH2PO4), and 10 gm of sodium chloride (NaCl), in 1000 ml of distilled water. pH is adjusted to 6.0+ 0.1. Each antigen pack usually consists of 10 ampoules of 5 ml buffered saline solution. Preparation of un buffered saline solution: This is prepared by adding 1 gm of sodium chloride to 100 ml of distilled water. III Specimen Serum and CSF.

PRINCIPLE PROCEDURE The VDRL slide flocculation test is a simple, rapid convenient and economical test for serodiagnosis of syphilis. This is an example of standard test of syphilis in which cardiolipin antigen is used to detect non-specific reaginic antibodies in the serum. Cardiolipin antigen is an alcoholic extract of beef heart tissue to which cholesterol and lecithin are added. This is a nontreponemal test because no treponemal antigen is used in this

Preparation of serum 1 Inactivate the serum in water bath at 56°C for 30 minutes. Note: After removal from the water bath, all the sera are examined and those found to contain particulate debris are recentrifuged. The quantity of serum used in each test is 0.05 ml.

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Preparation of antigen emulsion

Quantitative serum test

1 Pipette 0.4 ml of buffered saline to the bottom of a 1 oz reagent bottle with flat or concave inner bottom surface. 2 Add 0.5 ml of antigen, drawn from an ampoule in to a 1.0 ml pipette graduated to tip, directly on to the saline while rotating the bottle on a flat surface. The antigen is added drop by drop but rapidly so that it takes approximately 6 seconds to complete the delivery. 3 Blow the last drop of antigen and continue rotation of the bottle for 10 more seconds. 4 Then add 4.1 ml of buffered saline from a 5.0 pipette stopper the bottle and shake it vigorously for approximately 10 seconds. Note: Temperature of the buffered saline solutions and antigen should be preferably between 23°C to 29°C during the preparation of antigen emulsion.

Quantitative test is performed on all reactive serum samples and on all samples showing weakly reactive or rough in the qualitative test. Different dilutions of the serum are prepared in tubes as successive twofold dilutions (in the range of 1:2, 1:4, 1:8, 1:16, etc.) with buffered saline solutions. Each dilution of the serum is treated as an individual serum. These serum dilutions are tested as described above, in qualitative serum test.

Maturation of the antigen: It increases the sensitiveness and this is almost complete in 15 minutes to 30 minutes. The antigen emulsion prepared ought to be used during the same day. Preliminary testing of antigen emulsion: Each batch of prepared antigen emulsion should first be examined by testing known reactive and non-reactive sera. This test should present typically reactive and non-reactive results, and the number and distribution of antigen particles per microscopic field in the negative sera should be optimum.

Observe for the formation of floccules immediately after rotation under a microscope with low power objective (10 x magnification). The antigen particles are seen as small fusiform needles, which remain more or less evenly, disperse in a nonreactive serum, whereas these aggregate into clumps in reactive sera. Zone reactions are recognized by the irregular clumping, which are not compact. In such cases the results are reported on the basis of quantitative reaction done on the same serum.

Qualitative serum test

RESULTS AND INTERPRETATION

1 Pipette 0.05 ml inactivated serum into the cavity of a VDRL slide. Note: Slide must be thoroughly cleaned and then well dried before use. 2 One drop (1/60 ml) antigen emulsion is added on to the serum. The drop of antigen is transferred from an 18 gauze needle cut across and fitted to 1 ml syringe. 3 Rotate the slide for 4 minutes (180 rotations per minute). Note: If rotated by hand on a flat surface this movement should roughly circumscribe a 2 inch diameter circle 120 times per minute. 4 Read the result under 10 x objectives of the microscope.

1 Qualitative serum test

Quality control Known reactive (with known titre) and non-reactive sera are included.

OBSERVATIONS

The results are reported on the basis of quantitative reaction done on the same serum. No clumps or very slight roughness: Non-reactive. Small clumps: Weakly reactive. Medium and large clumps: Reactive.

2 Quantitative serum test Results are reported in terms of the highest dilution of the serum that produces a definite reaction.

Table 43-1 Advantages and disadvantages of VDRL test Advantages

Disadvantages

It is a simple and rapid diagnostic test. It helps in presumptive diagnosis of syphilis. It can be used as a prognostic test. It can be used as qualitative and quantitative tests. It is an inexpensive test. It is reproducible.

It is a non-specific test. It shows biological false positive reactions. This test alone is not confirmative. Improper temperature of the laboratory, specimens or reagents contributes to the error in the test.

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BOX 43-1 VDRL –ELISA An automated VDRL-ELISA test has been developed using cardiolipin antigen, which can measure IgG and IgM antibodies separately and is suitable for large scale testing of sera. The newest of the nontreponemal tests, VISUWELL Reagin, is based on the Pedersen method. In the indirect ELISA procedure, VDRL antigen coats the wells of a microtiter plate. This test has a sensitivity of 97% in untreated syphilis and specificity of 97%. The reactivity of this test disappears with treatment of the patient. The disadvantage of this test is the inability to quantitate the reactivity of a patient’s serum to an end point titre in order to assess the efficacy of treatment. Several hundred sera can be tested by VISUWELL Reagin test in a day.

BOX 43-2 BIOLOGICAL FALSE POSITIVE REACTIONS OF VDRL TEST Biological false positive (BFP) reactions are defined as positive reactions obtained in tests using cardiolipin antigen, with negative results in specific treponemal tests, in the absence of past or present treponemal infections, and not caused by technical faults. As cardiolipin antigen is present both in T. pallidum and in mammalian tissues, reaginic antibodies may be induced by treponemal or host tissue antigens. This accounts for BFP reactions. They represent the nontreponemal cardiolipin antibody responses. BFP reactions may occur in about one percent of normal sera. Clinically, BFP reactions may be classified as acute or chronic. Acute BFP reactions last only for a few weeks or months and are usually associated with acute infections, injuries or inflammatory conditions. Chronic BFP reactions persist for longer than six months and are typically seen in SLE and other collagen diseases. Leprosy, malaria, relapsing fever, infectious mononucleosis, hepatitis and tropical eosinophilia are examples of other conditions associated with BFP reactions.

KEY FACTS 1 2 3 4 5

VDRL is an example of slide flocculation test and is an example of standard test of syphilis. Cardiolipin antigen is used to detect non-specific reaginic antibodies in the serum. Maturation of the antigen increases sensitiveness of the test. Observe for the formation of floccules immediately after adding the antigen to the serum in a VDRL slide. Quantitative test is performed on all reactive serum samples and on all samples showing weakly reactive or rough in the qualitative test. 6 Zone reactions are recognized by the irregular clumping, which are not compact. 7 VDRL test shows biological false positives in a variety of conditions.

VIVA 1 2 3 4

What is the principle of the VDRL test? What is the antigen used in the VDRL test? What are the advantages and disadvantages of the VDRL test? List the biological false positive conditions shown by the VDRL test.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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44

Radial Immunodiffusion Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Quantitate the protein antigens in a test serum by single radial immuno diffusion.

Analytical balance, water bath, dark ground illuminating box, moist chamber, microscope slides, glass plates, beakers, syringes, micropipettes, Pasteur pipettes, measuring cylinder, measuring template and test tubes. III Specimen Serum.

INTRODUCTION Radial immunodiffusion is similar to the double diffusion method. In this test, antiserum is incorporated into the agar gel during its preparation. As in the double diffusion method, holes are cut in the agar, and antigen is placed in the well. As the antigen diffuses out of the well, it will form complexes with the incorporated antibodies, which are visualized as a ring. The size of the ring is proportional to the amount of antigen in the well. By constructing and using a standard curve, the amount of antigen in each well can be quantitated. Uses of gel diffusion tests are mentioned in the table 44-1.

PRINCIPLE When the radial diffusion of the soluble antigens from the cylindrical wells occurs in to the antibody incorporated gel, then circular precipitates develop. If the antibody is evenly distributed in the uniformly thick gel and well size and volume of the antigen is kept constant then the diameter of the precipitate is directly proportional to the antigenic concentration. This technique helps in the quantification of human serum proteins feasible even in a moderately equipped laboratory.

REQUIREMENTS I Equipments Electrophoresis power supply. II Reagents and glass wares Barbitone buffer, purified agar, Bacto agar, amido black, acetic acid and physiological saline.

PROCEDURE Preparation of antibody containing gels 1 Make 2% agar solution in barbitone buffer. 2 Boil the mixture till the gel particles are dissolved fully and allow it to cool down to 45-50°C . 3 Mix the appropriate quantity of the gel with the pre-warmed monospecific antibody (10% of the antibody is used). 4 Then mix and pour the gel on microscopic slide and allow setting.

Calibration of reference graph 1 A set of five standards (5, 6, 7, 10 and 15 md/dl) are selected and applied separately to the cylindrical wells using a suitable template. 2 Keep the gel in a moist chamber at 4°C . 3 Measure the diameters of the circular precipitates under oblique illumination after 24 hours by using the special measuring template. 4 Plot the graph between square of the diameters on the ordinate and the corresponding standard antigenic concentrations on abscissa using a linear graph paper. Note: A straight line is obtained by joining at least three reference points, which indicates accuracy of the graph. Alternatively, graph may be plotted between diameter of the ring and the antigenic concentration and in that case a curved line with a convexity upwards is obtained; a straight line or a line with concavity upwards indicates error in the technique.

Textbook of Practical Microbiology

Testing unknown serum samples 1 Dilute the test sample in a ratio of 1 in 15 with physiological saline. 2 Punch out a total of 9 wells (2mm wide) in the gel using the suitable template. Note: At least one check standard is employed on each occasion to test the reproducibility of the procedure while the remaining wells are charged with the test samples. 3 Transfer the gel to the moist chamber at 4°C. 4 Observe the gel after 24 hours under oblique illumination for ring precipitates. 5 Measure diameter of the ring and record it.

QUALITY CONTROL Known positive and negative samples.

OBSERVATIONS

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RESULTS AND INTERPRETATION 1 The antigen concentration is determined by taking the intercepts on the graph. The final value of the serum antigen level is derived by multiplying with the dilution factor (15). 2 The slide may be stained and preserved in a plastic bag for a permanent record. Table 44-1 Uses of gel diffusion tests 1 2 3 4

Screening sera for antibodies to influenza viruses. Diagnosis of small pox. Elek’s test for toxigenicity in diphtheria bacilli. Testing for normal and abnormal proteins in serum and in urine. 5 Detection of a Fetoprotein antigen in serum. 6 Detection of specific antigens of cryptococci and meningococci in cerebrospinal fluid. 7 Purification and identification of proteins and nucleic acids.

Observe the gels for precipitin lines after 24 hours under oblique illumination for ring precipitates.

KEY FACTS 1 2 3 4

Circular well with a clean edge is very important for a well-defined ring precipitate. Since there is a batch-to-batch variation the reference graph should be plotted for every new batch. The slope of the calibration graph is influenced by antibody concentration in the gel. The size of the ring is proportional to the amount of antigen in the well.

VIVA 1 What is the principle of gel diffusion test? 2 What are the advantages of gel diffusion? Ans. The advantages are as follows: a The reaction is visible as distinct band of precipitation. b Precipitin band is stable and can be stained for preservation. c The number of different antigens in the reacting mixture can be readily observed. 3 Classify gel diffusion methods? Ans. Immunodiffusion Single diffusion in one dimension (Oudin procedure). Double diffusion in one dimension (Oakley Fulthorpe procedure). Single diffusion in two dimensions (Radial immunodiffusion). Double diffusion in two dimensions (Ouchterlony procedure). Immunoelectrophoresis. Electroimmunodiffusion Countercurrentimmunoelectrophoresis. Rocket electrophoresis. 3 What are the uses of gel diffusion test? FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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45

Immunoelectrophoresis Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate different antigens in antigen mixtures by immunoelectrophoresis.

INTRODUCTION When mixtures, such as those in body fluids, are being analyzed in diffusion experiments like the double-diffusion system, it is difficult to distinguish the individual antigenic components from each other. Immunoelectrophoresis in principle is a method of combination of electrophoresis and diffusion. By these methods antigens present in a mixture are separated from each other by agar gel electrophoresis. Subsequently these antigenic components react specifically with their antibodies to form antigen-antibody precipitates that are formed in the gel. Immunoelectrophoresis is useful for demonstration of normal and abnormal serum proteins such as the myeloma proteins.

PRINCIPLE Immunoelectrophoresis techniques is carried out on a glass slide layered with semisolid gel. An aliquot of the antigen mixture is kept in the well cut out of the gel. Electrophoresis is carried out for one hour under electric current, this procedure permits separation of proteins. A rectangular trough is then cut parallel to the line of these separated proteins. Antiserum is placed in this trough. Diffusion is allowed to occur for 18-24 hours. The antiserum will diffuse laterally and react with the separated components. Individual precipitation lines develop with each separated components of the antigen mixture.

REQUIREMENTS I Equipments Electrophoresis power supply, and electrophoresis chamber.

II Reagents and glass wares Gel punch , glass plates, glass slides (3” x 2”) and standard laboratory wares. Buffer (pH 8.6), purified agar or agarose, amido black, acetic acid, paper wicks and physiological saline. III Specimen Patients serum. Human serum (antigen) and antihuman serum (antibodies)

PROCEDURE 1 Take a clean glass slide, To coat the slide place two to three drops of the molten agar and spread over the surface of the slide. 2 Put the coated slide on a horizontal work bench. 3 Pipette 6 ml of agar to the slide, spread it uniformly and allow it to set at room temperature, and then at 4°C in a refrigerator for 1 hour to set completely. 4 Remove the slide from the refrigerator. 5 Place the slide on a graph paper. Punch two wells at about one third of its length. Suck out the agar plugs from wells and discard. Note: These two wells should be 2 mm in diameter and I cm apart. 6 Fill the two wells with the serum. Note: the serum is mixed with the indicator dye (bromthymol blue)with a pipette. 7 Keep the slide in the electrophoresis chamber. Connect it to the buffer with Whatman 3 filter paper strips. Note: The slide should be kept in such a way that the well containing antigen will be near the cathode. This will make the components move toward the anode. 8 Fill the electrophoresis chamber with the buffer. 9 Connect it to the power supply and run the electrophoresis for 60-90 min at 10-15 mA/slide till the bromthymol blue (indicator dye) spreads to the end of the slide. 10 Remove the slides after turning off the power. Cut a trough between the two wells running parallel to the direction ofthe run.

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11 Remove the agar and fill the trough with the antiserum. 12 Keep the slide in the moist chamber and allow the diffusion to take place at 4 °C for 24 hours. 13 Remove the slides and observe for the lines of precipitation. 14 Wash the slide and stain the slide.

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OBSERVATIONS Observe the gels for precipitin lines after 24 hours at 4°C.

RESULTS AND INTERPRETATION QUALITY CONTROL Known positive and negative samples are used in the test.

1 The antigen components in the mixture is noted by observing different lines of precipitations. 2 The slide may be stained and preserved in a plastic bag for a permanent record.

KEY FACTS 1. Immunoelectrophoresis is useful for demonstration of normal and abnormal serum proteins such as the myeloma proteins. 2. Keep the slide in the electrophoresis chamber in such a way that the well containing antigen will be near the cathode. This will make the components move toward the anode. 3. Observe the gels for precipitin lines after 24 hours at 4°C.

VIVA 1 What is the principle of immunoelectrophoresis? 2 What are the advantages of immunoelectrophoresis? 3 List the applications of immunoelectrophoresis test.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

46

Counter-current Immunoelectrophoresis Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform counter-current immunoelectrophoresis (CIEP). 2 Demonstrate antibodies or antigens in serum and other body fluids by counter-current immunoelectrophoresis (CIEP).

INTRODUCTION Counter-current immunoelectrophoresis (CIEP) is a highly specific test used for demonstration of either antibodies or antigens in the serum, CSF, urine and other body fluids. This method was used earlier for diagnosis of many infectious diseases by demonstration of antigens or antibodies. The test is used for detection of hepatitis B antigen and antibodies in the serum, cryptococcal antigen in the CSF, hydatid antigens and antibodies in the serum, and amoebic antigens and antibodies in the serum. The CIEP is also used to detect hydatid and filarial antigens in the urine specimens. The CIEP test has the advantages of being simple, and rapid test, the result can be obtained within 30-45 minutes of performing the test. The test is also highly specific test. Disadvantages of the test is that it is a low sensitive test. The supporting medium, antiserum and its dilutions, control sera, buffer pH, voltage of the current, staining of the slides and other variables if not properly standardised may affect final result of the test.

PRINCIPLE The CIEP test is based on the movement of the antibodies toward the cathode while the antigens move in the opposite directions (toward the anode) in a layer of agarose on a glass slide under electric current. The test is carried out on glass slides layered with the agarose with the wells cut out off the agarose. One well is filled with antigen while the other well is filled with antibody. The slides are kept in an electrophoresis tank in such a way that the wells containing antigen will be on

the cathodic side and the well containing antibodies will be on the anodic side, so that during electrophoresis antibodies move towards the cathode and antigens move towards the anode. A line of precipitation visible to the naked eye is formed at a point between the antigen and antibodies. The test takes 30 minutes. The lines of precipitation in the slide can be stained by Amido black or 1% Coomassie brilliant blue stains. In this chapter the CIEP to detect hydatid antigen in the serum will be described.

REQUIREMENTS I Equipment Electrophoresis power supply and electrophoresis chamber. II Reagents and glass wares Glass plates, glass slides (7.5 cm by 2.5 cm), template, gel punch and standard laboratory wares. Veronal buffer, 0.075 M (pH 8.6), normal saline, purified agar or agarose, amido black,7% acetic acid and Whatman filter paper No.3. III Specimen Patients test serum, hydatid antigen, polyclonal hydatid antibodies raised in the rabbits Preparation of Veronal buffer 0.075 M (pH 8.6): The buffer is prepared by mixing dimethyl barbituric acid, 276 gm in 1000 ml of distilled water and dissolving it by heating. This is followed by adding 15.45 gm sodium dimethyl barbiturate and 1 gm of sodium azide to the solution. The pH is adjusted to 8.6. Preparation of agarose: This is prepared by adding 1gm agarose to 10 ml buffer and 90 ml distilled water. The agarose is melted by keeping it in a boiling water bath.

PROCEDURE 1 Take a clean glass slide, To coat the slide place two to three drops of the molten agar and spread over the surface of the slide.

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2 Put the coated slide on a horizontal work bench. 3 Pipette 2.5 ml of 1% agar to the slide, spread it uniformly and allow it to set at room temperature, and then at 4°C in a refrigerator for 1 hour to set completely. 4 Remove the slide from the refrigerator. 5 Place the slide on a template. Punch out parallel rows of wells 4mm in diameter on the slides at distance of 3 mm from each other Note: Nine pairs of wells are punched out on each slide. 6 Fill each well with 10µl of the patients serum to be tested and polyclonal hydatid antibodies. 7 Keep the slide in the electrophoresis chamber. Connect it to the buffer with Whatman 3 filter paper strips. Note: The slide should be kept in such a way that the well containing test sera will be near the cathode and the wells containing the hydatid antibodies will be near the anodic side. 8 Fill the electrophoresis chamber with the buffer. 9 Connect it to the power supply and run electrophoresis for 30 min at 10-15 mA/slide. 10 Remove the slides after turning of the power off. 11 Remove the slides and observe for the lines of precipitation. Note: The slides can be kept at 4 °C for 24 hr and the result read and stained. 12 To stain the slides wash the slides first in saline for several hours. Remove the salts by washing the slides several times in distilled water for one hour. 13 Stain the slides by immersing the slides in 1% Amido black in 7% acetic acid for 15-30 minutes. 14 Destain the slides with 7% acetic acid till the background stain is removed.

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15 After staining, the lines of precipitation stains dark blue.

QUALITY CONTROL Known hydatid antigen-positive and - negative serum samples. With every slide known hydatid antigen is placed in the wells on the cathodic side and hydatid antibodies in the wells on the anodic side as control.

OBSERVATIONS Observe the lines of precipitation after 30 min. First observe for the lines of precipitation in between the wells containing known hydatid antigen and known hydatid antibodies (Control row). Then observe for the lines of precipitation in the test sera.

RESULTS AND INTERPRETATION 1 The line of precipitation is present in the control row : The test is performed correctly. 2 Line of precipitation is formed between the test serum and hydatid antibodies wells: It indicates the patient serum is positive for hydatid antigen by the CIEP test. 3 No line of precipitation is formed between the test serum and hydatid antibodies wells: It indicates the patient serum is negative for hydatid antigen by the CIEP test.

KEY FACTS 1 The CIEP test is based on the movement of the antibodies toward the cathode while the antigens move in the opposite directions (toward the anode) in a layer of agarose on a glass slide under electric current. 2 The CIEP test has the advantages of being simple, and rapid test, the result can be obtained within 30-45 minutes of performing the test. 3 After staining by Amido black, the lines of precipitation stains dark blue.

VIVA 1 2 3 4 5

What is the principle of counter current electrophoresis CIEP test? What are the advantages of CIEP test? What are the disadvantages of CIEP test? What are the dyes used for staining the slide after electrophoresis? List the applications of CIEP test.

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

47

Indirect Haemagglutination Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform indirect haemagglutination test. 2 Demonstrate antibodies in serum by the indirect haemagglutination (IHA) test.

INTRODUCTION Indirect haemagglutination (IHA) test is a technically simple and inexpensive technique using antigen sensitized erythrocytes for demonstration of serum antibodies in a variety of parasitic diseases. The test has been proved useful for diagnosis of various parasitic diseases viz., schistosomiasis, echinococcosis, leishmaniasis, malaria amoebic liver abscesses. Erythrocytes from different sources have been used by various workers in the IHA test. Frequently and widely used erythrocytes are from sheep. Other sources are from human of ‘O’ blood group, turkey, goose, bovine and swine Avian cells are usually recommended because being nucleated, they are heavier, they settle quickly, so results of hemagglutinated chick cells could be observed within 30-45 min of agglutination. The IHA using the chick cells have been used in the IHA for demonstration of specific serum antibodies in the diagnosis of amoebiasis, echinococcosis and filariasis. Reverse passive haemagglutination test is a different test used to detect antigen in the serum and other body fluids (Table 47-1). In this chapter the IHA method for demonstration of hydatid antibodies in the serum for diagnosis of hydatid disease will be described.

PRINCIPLE This is based on the principle that the red blood cells act as carrier particles for the antigen (against which antibodies will be demonstrated). The RBCs sensitized with antigen are added

to the patients sera to demonstrate specific antibodies. The haemagglutination pattern of the antigen sensitized-RBCs with the sera are noted. Formation of button with test sera indicate that agglutination of RBCS has not occurred. Matt formation with test sera indicates that agglutination of RBCS has taken place. In IHA for hydatid disease, the chick RBCs are sensitized with hydatid antigen (fluid obtained from human hydatid cyst). The haemagglutination pattern of chick RBCs are observed after 30-45 min of the test.

REQUIREMENTS I Lab wares Microtitre plates, droppers (25µl) and diluters (25µl). II Reagents Hydatid antigen, double aldehyde stabilized chick RBCs, phosphate buffer saline (pH7.2), phosphate buffer saline (pH6.4) and diluent (PBS pH 7.2 with 0.1% bovine serum albumin). Chick RBCs are stabilized by double aldehyde stabilization method. Preparation of hydatid antigen: Hydatid cysts removed surgically from humans or animals (cattle) are the source of hydatid fluid. The intact cyst should be collected as such without any rupture and sent to the laboratory immediately. The hydatid cyst should not be preserved in the formalin. The fluid is aspirated aseptically by syringe from an intact hydatid cyst. This hydatid fluid is checked microscopically for the presence of any scolices or hooklets. The fluid obtained is sterilized by method of filtration using a Seitz filter or a membrane filter. The fluid is then checked for sterility by inoculating in a blood agar and McConkey agar and incubating aerobically. The protein content of the fluid is estimated by autoanalyser. The antigens is stored in -20ºC in 1ml aliquots. The optimum sensitizing dose (OSD) of the antigen is determined for each dilution of stabilized chick cells by chequer board titration against the known positive control and negative

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control. The lowest amount of antigen which shows the maximum heamagglutination with the known positive control and negative reaction with the negative control is taken as OSD of the antigen for that batch. III Specimen Serum, positive control serum and negative control serum.

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QUALITY CONTROL Known hydatid antibody-positive and - negative serum samples. With every test, known hydatid antibody positive and negative serum samples are tested in the microtiter plate. A well containing only sensitized chick cells (no sera is added to this well) is used as cell control.

PROCEDURE Sensitisation of chick RBCs with OSD of the antigen. 1 Take 0.1ml of packed DAS chick cells in a centrifuge tube and wash with PBS 7.2 (twice) and centrifuge. 2 Wash with PBS 6.4 and centrifuge. 3 Add 0.1ml of this packed chick cells to 0.9ml of optimum sensitizing dose (OSD) of the hydatid antigen. 4 Keep the chick cells in water bath for 5 minutes at 50°C with intermittent shaking at the end of each minute. 5 Keep the cells at 4°C overnight. 6 Next day mix the cells and keep the cells again in waterbath at 50°C for 10 minutes. 7 Wash twice with PBS 7.2 and once with 0.1% bovine albumin (prepared in PBS 7.2). 8 Make 1% suspension with 0.1% BSA. 9 Store it at 4 °C until use.

Performance of the IHA test 1 Inactivate the test serum at 56ºC for 30 minutes before starting the test. 2 Dispense a 25µl volume of the diluent (PBS pH 7.2 with 1% bovine serum albumin) in each well of U bottomed microtitre tray. 3 Add 25µl volume of the serum to the first well of the appropriated row. 4 Dilute the sera upto the eleventh well leaving the last well as a serum free control. 5 Add 25µl volume of the 1% chick cells sensitized with the OSD antigen to each well. 6 Agitate the plate gently for 2 min. 7 Incubate the plates at room temperature for 30 min and note the pattern of haemagglutination.

OBSERVATIONS Formation of button in the cells with test sera indicate that agglutination of RBCs has not occurred Matt formation with test sera indicates that agglutination of RBCs has taken place. Matt formation in the cell control well rules out non-specific agglutination of RBCs.

RESULTS AND INTERPRETATION Formation of button in the cells with test sera indicate that agglutination of RBCs has not occurred. The serum showing a titre of 1:128 and below is considered negative for hydatid disease. Matt formation with test sera indicates that agglutination of RBCs has taken place. The serum showing a titre of 1: 128 and above is considered positive for hydatid disease. Matt formation in the cell control well rules out non-specific agglutination of RBCs.

BOX 47-1 REVERSE PASSIVE HAEMAGGLUTINATION TEST In this test, RBCs act as carrier particles for polyclonal and monoclonal antibodies for demonstration of specific antigen in the serum, CSF and other body fluids. Reverse passive haemagglutination test (RPHA) is used for demonstration of specific antigens for diagnosis of Hepatitis B infection, Japanese encephalitis, tuberculous meningitis and many other infectious diseases.

KEY FACTS 1 2 3 4 5

The red blood cells from different sources have been used by various workers in the IHA test. The red blood cells act as carrier particles for the antigen (against which antibodies will be demonstrated). The RBCs sensitized with antigen are added to the patients sera to demonstrate specific antibodies in serum. The IHA test is carried out in a microtiter plate. The lowest amount of antigen which shows the maximum haemagglutination with the known positive control and negative reaction with the negative control is taken as OSD of the antigen for that batch. 6 The serum showing a titre of 1: 128 and above is considered positive for hydatid disease.

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VIVA 1 2 3 4

What is the principle of IHA test? List different RBCs which can be used in the IHA test. List the uses of IHA test. What is reverse passive haemagglutination test?

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

48

Immunofluorescence Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate bacterial antigen by immunofluorescence test.

INTRODUCTION Immunofluorescence assays involve the use of either antigen or antibody labeled with a fluorescent substance. The fluorescent dyes have the property of absorbing the light of one wavelength (e.g., ultra violet light) and reflect back the light of a different wavelength (visible light). These reflected lights are visualized by a fluorescent microscope under ultra violet radiation.. Fluorescein isothiocyanate (FITC) is the most common dye used in the test. The dye emits a greenish/yellow light. Rhodamine (red/orange), dansyl (yellow), and phycoerythrin are the other dyes used in the test. Direct fluorescent antibody test is employed to detect the specific antigens of bacteria, viruses, parasites or other antigens in the serum, CSF, urine, faeces, tissues and other body fluids. This test is most frequently used to detect rabies virus antigen in the tissue collected from the skin on the nape of the neck or face. This is also used to detect Corynebacterium diphtheriae, Neisseria gonorrhoeae, measles and mumps in various clinical specimens.

PRINCIPLE In this test specific antibodies raised against the antigen to be detected (e.g., anti-rabies antibodies to detect rabies antigen) is labeled with a fluorescent dye, and is used to detect unknown antigen (e.g., rabies antigen) in the specimen. If antigen is present in the specimen, the same will bind the fluorescein labeled antibodies, and the antibody-bound fluorescein will emit a fluorescence, which will be observed by a fluorescent microscope using ultra violet radiations.

In this experiment you will use the method to demonstrate unknown bacterial antigens. The fluorescein labeled antibody is tagged to alcohol or acetone fixed bacterial smear. This is followed by washing the smear with physiologically buffered saline. During this step any uncombined fluorescent antibody will be washed away. In a positive test, the final reaction will be observed by a green fluorescence when observed under a fluorescent microscope. Advantages and disadvantages of the immunofluorescence test are summarized in the table 48-1.

REQUIREMENTS I Equipments Fluorescent microscope. II Reagents and lab wares Acetone, phosphate buffered saline (pH 7.2), buffered glycerol (pH 7.2): 10% glycerol in PBS, Evan’s blue counter stain (1:10000), and bovine serum albumin. Glass slides, Petri dishes, U-shaped glass rods to fit into petri dishes, filter paper, and Coplin jar and glass marking pencil. Commercially available fluorescent antibody Streptococcus group A and fluorescent antibody Enterococcus group D. III Specimen Broth cultures of Group A Streptococcus pyogenes and Group D Enterococcus faecalis incubated at 35°C for 24 hours. Unknown mixed broth cultures of Group A S. pyogenes/ Escherichia coli and Group D E.faecalis / E. Coli

PROCEDURE 1 Take three clean glass slides. 2 With glass marking pencil label first slide as S. pyogenes, and second slide as E. faecalis. Divide the third slide in half and label as mixed unknowns.

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3 Prepare alcohol or acetone fixed smears of S. pyogenes on the first slide, of E. faecalis on the second slide and of mixed unknowns on the third slide. 4 Add one drop of fluorescent antibody Streptococcus group A on to the slide one and fluorescent antibody Enterococcus group D to the second slide and spread gently over the surface of the smear. 5 In the third slide, label one side FA-A and the other slide FA-D and add one drop of fluorescent antibody Streptococcus group A to the side FA-A and one drop of fluorescent antibody Enterococcus group D to the side FA-D. 6 Allow to spread gently over the surface of the smear. 7 Place the prepared slides on the U-shaped glass rod kept in a petri dish and incubate at 25°C for 30 minutes. 8 Remove the slides from the petri dish and wash with 1% buffered saline to remove away excess antibody. 9 Put the slides in a Coplin jar containing 1% buffered saline for 10 minutes at 25°C. 10 Blot dries the slides with paper. 11 Add one drop of buffered glycerol to each slide and cover with a cover slip. Note: 1-10 µg/ml of phenylene diamine could be added to the mounting medium to prevent the fading of fluorescence of fluorescein. 12 Examine under a fluorescent microscope.

QUALITY CONTROL The slide with S. pyogenes and E. faecalis treated with fluorescent antibody Streptococcus group A and fluorescent antibody Enterococcus group D are used as known positive controls.

OBSERVATIONS The slides are observed for the absence or presence of fluorescence in all the slides. First slide with S. pyogenes : Second slide with E. faecalis : Third slide, the side labeled FA-A : Third slide, the side labeled FA-D :

Positive for fluorescence. Positive for fluorescence. Negative for fluorescence. Positive for fluorescence.

RESULTS AND INTERPRETATION The unknown specimen contains the bacteria E. faecalis but does not contain S. pyogenes.

Table 48-1 Advantages and disadvantages of the immunofluorescence test Advantages

Disadvantages

More sensitive. More specific when fluorescent dye is labeled to monoclonal antibodies. It can avoid the danger of radiation-based hazards. Rapid method. Useful for the identification of viruses.

Difficult to quantitate the test. Requires UV radiation for visualization of the test result. Requires expertise personnel. Fluorescent dye fades faster. Improper washing steps cause a very high background. False positive reactions may occur. Requires expensive equipment.

KEY FACTS 1 2 3 4 5

Immunofluorescence assays involve the use of either antigen or antibody labeled with a fluorescent substance. Direct fluorescent antibody test is employed to detect the specific antigens. Slides should be cleaned before use. The smears should not be allowed to dry at any stage. To minimize non-specific binding of proteins to cells or tubes, all dilutions and washings should be done in PBS containing 0.1 – 1% BSA. 6 1-10 µg/ml of phenylene diamine could be added to the mounting medium to prevent the fading of fluorescence of fluorescein.

VIVA 1 2 3 4

What is the principle of the direct fluorescent antibody test? What are the uses of direct fluorescent antibody test? Give examples of fluorescent dyes. What are the advantages and disadvantages of the direct fluorescent antibody test?

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FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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LESSON

49

Enzyme-linked Immunosorbent Assay

LEARNING OBJECTIVES

PRINCIPLE

After completing this practical you will be able to:

1. The sandwich ELISA for antigen: This method to detect antigen is very sensitive. The walls of the microtiter plates are coated with specific antibody against the antigen to be detected. The specimen such as serum to be tested are added to the coated wells. If antigen is present in the serum, then it will combine with the coated antibody. Antibody conjugated with enzyme is added to detect the antigen-antibody reactions. The conjugated antibody combines with antibody that has combined with the antigen. A substrate is added to detect the binding of conjugated antibody to antigen-antibody complex. In a positive test, the positive reaction is tested by the change of colour, which can be read by spectrophotometer or ELISA reader. 2. The indirect ELISA for antibodies: This is a frequently used method to detect antibodies. Polystyrene tubes or wells in polystyrene plates are coated with the antigen solution. An aliquot of the serum to be tested is added to each tube and is incubated, followed by washing. To detect the reaction a goat antihuman immunoglobulin antibody conjugated with enzyme is added. The enzyme conjugated antihuman immunoglobulin binds to the antibody. A substrate is added to detect the binding and in a positive reaction the enzyme acts on the substrate to produce a change of colour. Then the enzyme-linked antiglobulin is added. If antibodies are present in the serum, they will have bound to the antigen, which has bound to the plate, and now the antiglobulin will bind to the primary antibodies. Add the substrate for the enzyme. If the reaction occurs (for example, if the solution changes color), this means that the enzyme-linked antiglobulin was bound, which means that the serum did, in fact, contain the antibodies of interest. The color change can be quantified using a spectrophotometer. 3. Competitive ELISA for antibodies: This involves the use of two specific antibodies. One antibody is conjugated with enzyme; where as the other antibody is present in the serum to be tested. Competition occurs between these two antibodies for the same antigen, hence is called the competitive ELISA. This is used for demonstration of HIV antibodies in the serum.

1 Know the importance of various steps involved in performing enzyme-linked immunosorbent assay (ELISA). 2 Perform the ELISA test and interpret the results.

INTRODUCTION Enzyme-linked immunosorbent assay (ELISA) is a widely used method for demonstration of specific antigens or antibodies in the serum and other body fluids. They are used extensively for diagnosis of a variety of human diseases including many viral (AIDS, hepatitis, rubella, respiratory syncytial infection, etc.), parasitic (amoebiasis, cysticercosis, leishmaniasis, lymphatic filariasis, etc.), bacterial (syphilis, brucellosis, salmonellosis, etc.) and fungal (histoplasmosis, aspergillosis, etc.). The principle of ELISA in essence is similar to that of the fluorescence, the only difference being that an enzyme is used instead of fluorescent dye. The most commonly used enzymes are alkaline phosphatase, horseradish peroxidase and ßgalactosidase. Enzyme activity is generally determined by the change of colour, which can be read by spectrophotometer or ELISA reader. ELISA is a heterogenous enzyme assay that uses solid phases e.g. plastic tubes, polyvinyl chloride plates, beads, microplate, and membranes such as cellulose acetate or cellulose nitrate. The enzyme immunoassay can be broadly of three types: a. the sandwich ELISA for antigen, b. the indirect ELISA for antibodies, and c. competitive ELISA for antibodies. Depending on the nature of the antigen used, the ELISA can also be classified as 1st, 2nd and 3rd generation ELISA as used in HIV infections (Box 49-1). Enzymes and substrates used in the ELISA are listed in the table 49-1.

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The Dot ELISA is a rapid visually read microassay form of the ELISA (Box 49-2). The advantages and disadvantages of ELISA tests are mentioned in the table 49-2. In this experiment indirect ELISA to detect antibodies and sandwich ELISA to detect antigen will be described.

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7 Add 100 µl of conjugate to each well and incubate at 37°C for 1 hour. 8 Wash the wells thrice and add 100 µl of substrate solution and incubate for 30 minutes at 37°C in dark. 9 Stop the reaction with 50µl of 3N sulphuric acid (H2SO4) to each well. 10 Read the results with naked eye or at 492 nm in an ELISA reader.

INDIRECT ELISA TO DETECT ANTIBODIES QUALITY CONTROL REQUIREMENTS I Equipments Micro titer plate, ELISA reader and ELISA washer.

II Reagents and glass wares Carbonate buffer (pH 9.6), washing buffer (PBS-Tween 20), conjugate, citric acid phosphate buffer (0.1M) pH 5.0, substrate, 4% bovine serum albumin (BSA) in PBS-tween 20 and 3N H2SO4, micro pipettes, tips, glass beakers, and filter paper. 1 Carbonate buffer (pH 9.6) Solution A: 5.3 g sodium carbonate (Na2CO3) is dissolved in one liter of double distilled water. Solution B: 1.2 g sodium bicarbonate (NaHCO3) is dissolved in one litre of double distilled water. Mix 16 ml of solution A with 34 ml of solution B and make volume to 100 ml with distilled water and adjust pH to 9.6. 2 Washing buffer (PBS-Tween 20): Phosphate buffered saline (0.01M) pH 7.2 with 0.05% tween 20. 3 Conjugate: Goat antimouse immunoglobulin conjugated with horse radish peroxidase. 4 Citric acid phosphate buffer (0.1M) pH 5.0: Weigh and dissolve 7.3 g of citric acid and 11.86 g of disodium hydrogen phosphate (Na2HPO4) in one liter double distilled water and adjust pH to 5.0. 5 Substrate: Dissolve 8 mg of Orthophenylene diamine in 15 ml of citrate buffer. Then add and mix 15 ml of Hydrogen peroxide just before to use.

III Specimen Serum, CSF, tear, etc.

PROCEDURE 1 Make up antigen to optimal dilution in carbonate buffer and add 100 µl per well in 96 well microtiter plate and incubate at 37°C for 3 hours or at 4°C overnight. 2 Decant the antigen solution and wash the plates thrice with washing buffer. Remove all residual fluid. 3 Add 100 µl of 4% BSA in washing buffer to each well and incubate at 37°C for 1 hour. 4 Wash thrice with wash buffer and remove all residual fluid. 5 Add 100 ul of diluted serum samples and incubate for 1 hour at 37°C . 6 Wash the wells with washing buffer and remove all residual fluid.

Appropriate controls, such as controls with positive and negative sera, buffer control, conjugate control and substrate control should be tested along with every plate.

OBSERVATIONS Observe the microtiter plate for the development of colour. Observe control (negative, positive and blank) wells.

RESULTS AND INTERPRETATION Development of yellow colour indicates the test serum contains antibodies to tested antigen. No colour is negative reaction (Fig. 49-1).

SANDWICH ELISA TO DETECT ANTIGEN REQUIREMENTS I Equipments Micro titer plate, ELISA reader and ELISA washer.

II Reagents and glass wares Carbonate buffer, blocking solution, washing buffer, monoclonal antibodies (1-10 µg/ml in carbonate buffer) and substrate solution, micropipettes, tips, glass beakers, and filter paper.

III Specimen Serum, CSF, urine etc.

PROCEDURE 1 Coat the ELISA plates with monoclonal antibody (mAb) in carbonate buffer by adding 100µl to each well, and incubate overnight at 4°C . 2 Remove monoclonal antibody from ELISA plate and add 100µl of blocking solution. Incubate for 30 minutes at 37°C. Wash the plates thrice with washing buffer. Remove all residual fluid. 3 Add 100µl of 4% BSA in washing buffer to each well and incubate at 37°C for 1 hour.

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Enzyme-linked Immunosorbent Assay

4 Wash thrice with wash buffer and remove all residual fluid. 5 Add 100µl of diluted serum samples and incubate for 1 hour at 37°C. 6 Wash the wells with washing buffer and remove all residual fluid. 7 Add 100µl of conjugate to each well and incubate at 37°C for 1 hour. 8 Wash the wells thrice and add 100µl of substrate solution and incubate for 30 minutes at 37°C in dark. 9 Stop the reaction with 50µl of 3N H2S04 to each well. 10 Read the results with naked eye or at 492 nm in an ELISA reader.

RESULTS AND INTERPRETATIONS Development of yellow colour indicates the test serum contains antigens to tested antibody. Development of no colour inducates negative reaction.

QUALITY CONTROL Appropriate controls, such as controls with positive and negative sera, buffer control, conjugate control and substrate control should be tested along with every plate.

OBSERVATIONS Observe the microtiter plate for the development of colour. Observe control (negative, positive and blank) wells.

FIGURE 49-1 ELISA test.

BOX 49-1 1ST, 2ND AND 3RD GENERATION ELISA 1st generation ELISA: It is an ELISA in which crude antigen is used for the detection of antibodies. 2nd generation ELISA: It is an ELISA in which semi-purified antigen is used for the detection of antibodies. 3rd generation ELISA: It is an ELISA in which recombinant antigens or synthetic peptides are used for the detection of antibodies.

BOX 49-2 DOT ELISA Dot ELISA is a rapid visually read microassay. The principle is similar to that of ELISA where enzyme is used as a marker or label to detect the binding of antigen and antibody. The test is performed on a nitrocellulose membrane. Enzyme converts colourless substrate to a coloured product, which indicates the presence of antigen-antibody binding. When the test serum is layered on to the membrane, specific antibodies if present, will bind to corresponding dot of the antigen. Addition of a labeled serum antibody (conjugate) and the subsequent development of the colour allow the detection of the presence of antibodies based on the pattern of the antigen sites. Dot-ELISA can be used for the detection of antibodies as well as antigen. Table49-1 Enzymes and substrates used in the ELISA Enzymes

Substrates

Horseradish peroxidase. Alkaline phosphatase. b-D galactosidase, Glucose 6-phosphate dehydrogenase (G6PD). Acetylcholinesterase.

Hydrogen peroxide (H2O2). Nicotinamide adenine dinucleotide phosphate (NADP+). 2-nitrophenyl-b-D-galactoside. Glucose-6 phosphate. Acetylcholine.

Table 49-2 Advantages and disadvantages of ELISA test Advantages

Disadvantages

Highly sensitive Specific Rapid

Requires specialized expert personnel. Requires expressive laboratory equipment. Enzymes require cold environment for storage.

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KEY FACTS 1 2 3 4

Preliminary chequer board titrations are carried out for standardizing antigen and antibody concentration. Read all plates at a standard time after addition of the substrate because colour is not stable. Substrate should be made just before use as it decomposes spontaneously and is unstable. OPD should be stored at 4°C in the dark. It is carcinogenic.

VIVA 1 Expand ELISA. Ans. Enzyme-linked immunosorbent assay. 2 What is the principle of enzyme immuno assay? 3 What are different steps in the ELISA procedure? Ans. a Coating solid surface with antigen/antibody. b Binding antibody/antigen from test sample. c Binding conjugate. d Addition of substrate. e Observe the colour change and read the O.D. values. 4 What are various types of the ELISA? Ans. Competitive ELISA, sandwich ELISA, indirect ELISA, membrane bound ELISA, cassette ELISA, biotin-avidin ELISA, protein-A ELISA, etc. 5 What are the enzymes and substrates used in the ELISA? 6 What do you understand by the first, second and third generation ELISA? 7 What are the advantages and disadvantages of the ELISA?

FURTHER READINGS 1 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworth, London, 94-96, 1995. 2 Koneman EW, Allen SD, Janda WM, Schreckenbergu PC and Winn Jr. WC. Color Atlas and Textbook of Diagnostic Microbiology. 5th Edition. Lippincott Williams and Wilkins. 1997; pp. 1395. 3 Stites DP. Terr AI and Parslow TG. Medical Immunology. 10th Edition. 2001. pp. 902.

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UNIT

VII Microbial Genetics and Molecular Techniques

Introduction Lesson 50 Isolation of Plasmids Lesson 51 Polyacrylamide Gel Electrophoresis Lesson 52 Isolation of Antibiotic Resistant Mutant Lesson 53 Bacterial Conjugation

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Introduction The objectives of the study are to demonstrate the genetic methods such as isolation of plasmids, polyacrylamide gel electrophoresis (PAGE), isolation of antibiotic resistant mutants and conjugation in bacterial systems. Genetic information is stored in DNA of the bacteria as a sequence of nucleotide bases (adenine, cytosine, guanine, thymine, abbreviated as A, C, G, T respectively) read sequentially in a 5' to 3' direction (or in RNA, with uracil, abbreviated U, replacing thymine). The most common form of DNA (present in all cellular genomes, as well as many viral genomes) is double stranded. The 5' to 3' polarity of the two strands is opposite, and they are held together by hydrogen bonding between nucleotide base pairs, A to T and G to C. The sense strand carries the coded genetic information. The antisense strand consists of a complementary sequence of bases oriented in the opposite 5' to 3' direction. During DNA replication, the two strands separate and each is used as a template for synthesis of a new complementary strand. This allows genetic information to be replicated with a high level of precision. Because replication is bidirectional, new DNA can only be synthesized in a 5' to 3' direction, the overall pattern of replication is rather complex. Genetic information is transcribed from DNA to RNA, with the antisense strand of the DNA serving as a template for synthesis of RNA with the same base sequence (5' to 3') as the sense strand of the double helical DNA, except that uracil (U) replaces thymine (T). Genetic information contained in messenger RNA (mRNA) is translated into a sequence of amino acids in a polypeptide chain during protein synthesis (translation). A redundant nucleotide triplet code, read 5' to 3' on the mRNA (and on the sense strand of the DNA), specifies the amino acid sequence of the protein, read from N-terminal to C-terminal. The study of genetic materials helps to a). demonstrate mutations, b). study the gene transmission among progeny or generations, c) treat genetic disorders, and d) in diagnosis and research.

FREQUENTLY USED TERMINOLOGY IN GENETICS Phenotype: Phenotype refers to “the observable outward appearance of an organism, which is controlled by the genotype and its interaction with the environment”. Genotype: Genotype refers to “the genetic makeup of an individual organism”. Transcription: The “central dogma” of molecular biology describes the transcription of genetic information from a DNA nucleotide triplet code to an RNA triplet code, followed by translation to specific amino acid sequences in protein. Prototrophs: A wild type strain that has minimal requirements for exogenously supplied nutrients is referred to as a prototroph. Auxotrophs: A mutant strain that has lost the ability to synthesize its own supply of a particular nutrient, such as histidine or adenine or thiamine is called an auxotroph. Recombinant DNA technology: Recombinant DNA in this context refers to the creation of a new combination of DNA segments that are not found together naturally. Such technology is now widely used in many practical applications ranging from basic research on control of gene expression to forensic medicine to biotechnology. Restriction endonucleases: An endonuclease is an enzyme that can cleave the phosphodiester bonds of a nucleic acid at an internal site (as opposed to cleavage by an exonuclease, which can only remove nucleotides from one of the ends of a nucleic acid). Example: Eco RI G|AATTC. Hybridization probes: Complementary strands of DNA, RNA, or DNA plus RNA hybridize readily to form double stranded helical structures when placed under suitable annealing conditions. This property is used extensively in molecular genetics to identify specific nucleic acid sequences. A probe consisting of radioactively labeled DNA (or RNA) is hybridized to denatured DNA (or naturally single-stranded RNA) immobilized on a support, such as a nitrocellulose membrane. Hybridization is normally done at a temperature about 25°C below the melting (denaturation) temperature for the DNA.

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LESSON

50

Isolation of Plasmids

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Isolate the plasmid from bacteria.

INTRODUCTION Bacterial plasmids are small, circular, extra chromosomal DNAs that are capable of autonomous replication within bacterial cells. Plasmids have their own origins of replication, such that they can multiply independently within the bacterial cells. They usually contain only a few thousand base pairs of DNA and carry only a few genes, often for antibiotic resistance. Plasmids that carry appropriate genes are capable of making bacteria resistant to antibiotics(Box 50-1), which makes it possible to select for bacteria that have taken up the plasmids.

PRINCIPLE This method utilizes the molecular characteristics of covalently closed circular deoxyribonucleic acid (DNA) that is released from cells under conditions that denature DNA by using alkaline sodium dodecyl sulfate. Under these conditions, chromosomal DNA concentrations are reduced or eliminated. The DNA is freed of RNA and proteins by RNase and protease treatments. Just before extraction, the plasmid DNA is released from the folded chromosomal complex by a shearing step or by RNase treatment. The plasmid DNA can be resolved as covalently closed circular molecules by centrifugation in ethanol containing ethidium bromide. The clarified extract is used directly for electrophoretic analysis.

REQUIREMENTS I Equipments Centrifuge and inoculating chamber.

II Reagents and lab wares Ethidium bromide solution, Luria-Bertani medium, micropipettes, Eppendorf tubes, and Bunsen burner. Preparation of ethidium bromide stock solution: The solution is prepared by dissolving 10 mg of ethidium bromide powder in 1 ml of double distilled water. Preparation of ethidium bromide working solution: The solution is prepared bu adding 10µl ethidium bromide stock solution in 100 ml of agarose solution. Preparation of Luria-Bertani medium (LB medium): The medium is prepared by adding the following ingredients: Bactotryptone, 10 g; Bacto yeast extract, 5 g; sodium chloride (NaCl), 10 g; and distilled water, 1000 ml. The pH is adjusted to 7.0. III Specimen Bacterial strain: Plasmid containing strain PHSV 106 or equivalent strain is grown in a medium containing 20µg/ml ampicillin. The medium contains the following solutions: Solution A 50 mM glucose. 25 mM Tris chloride (ph 8). 10mM EDTA (pH 8). Solution B 0.2 N sodium hydroxide (NaOH). 1% SDS. Solution C 5M potassium acetate, 60 ml. Glacial acetic acid, 11.5 ml. Distilled water, 28.5 ml.

PROCEDURE Extraction of plasmid 1 Inoculate a single bacterial colony into 2 ml of LB medium containing antibiotic ampicillin in a 15 ml test tube and incubate over night with vigorous shaking.

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2 Take 1.5 ml of culture into a Microfuge tube and centrifuge at 12, 000 g for 2 min at 4°C and discard the supernatant. 3 Resuspend the pellet in 100 µl of ice-cold solution A and vortex vigorously. 4 Add 200 µl of solution B and mix the contents by inverting the tube rapidly several times. 5 Add 150 µl of ice-cold solution C and mix the contents by inverting the tube. 6 Store the tube in ice for 5 minutes. 7 Centrifuge at 12, 000 g for 5 minutes at 4°C and transfer the supernatant to a fresh tube. 8 Precipitate the DNA with two volumes of 100% ethanol at room temperature. Mix well and allow it to stand for 2 minutes. 9 Centrifuge at 12000 g for 5 minutes at 4°C and remove supernatant. 10 Resuspend the pellet in 50 µl of TE (pH 8) containing RNAse (20 µg/ml). 11 Run in agarose gel containing ethidium bromide and view under UV transilluminator. Electrophoresis on agarose gel 1 Prepare 1% agarose in 1 X TE buffer, melt in a boiling water bath and add 1 µl of ethidium bromide. 2 Pour 20 ml -25 ml of molten agarose in the gel chamber after placing the comb and allow it to set. 3 Place the gel in electrophoresis tank, remove the comb and pour the TE buffer until it fully covers the gel surface. 4 Load the wells with 5 µl plasmid preparation and 2 µl bromophenol blue tracking dye and a molecular weight marker in other well. 5 Run electrophoresis at 50-70 V/gel till the tracking dye reaches the end.

6 Remove the gel carefully and place on UV transilluminator and examine for the bands.

QUALITY CONTROL A parallel procedure should be carried out using standard strain of bacteria. Molecular weight marker must be used in the analysis.

OBSERVATIONS Observe the stained gel under UV illumination for DNA band. Observe the bands of molecular weight marker.

RESULTS AND INTERPRETATION Plasmid appears as a sharp orange band. Undigested RNA appears as hazy opaque area along the lanes.

BOX 50-1 ROLE OF PLASMIDS IN DRUG RESISTANCE Bacterial plasmids, covalently closed circular DNAs, have their own origins of replication and are capable of autonomous replication. They usually contain only a few thousand base pairs of DNA and carry only a few genes, often for antibiotic resistance. Plasmids that carry appropriate genes are capable of making bacteria resistant to antibiotics, which makes it possible to select for bacteria that have taken up the plasmids.

KEY FACTS 1 Plasmid is small, circular, extra chromosomal DNA that is capable of autonomous replication within bacterial cells. 2 Care must be taken at every step of the procedure for extraction of the plasmid DNA and electrophoresis on agarose gel. 3 Care must be taken while decanting the supernatant after centrifugation.

VIVA 1 What are plasmids? 2 What are the properties and functions of plasmids? Ans. Plasmids code for synthesis of a few proteins not coded for by the nucleoid. For example, R-plasmids, found in some Gram-negative bacteria, often have genes coding for both production of a conjugation pilus and multiple antibiotic resistance. Through a process called conjugation, the conjugation pilus enables the bacterium to transfer a copy of the R-plasmids to other bacteria, making them also multiple antibiotics resistant and able to produce a conjugation pilus. In addition, some exotoxins, such as the tetanus exotoxin and Escherichia coli enterotoxin are also coded for by plasmids. 3 How do you extract plasmids from bacteria?

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FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Systemic Bacteriology. Volume 2.. Arnold publishers. pp. 2002, pp 1501. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. pp 94-96. 3 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. pp. 921.

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51

Polyacrylamide Gel Electrophoresis

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to: 1 Prepare agarose gel. 2 Assemble the electrophoresis apparatus and power pack. 3 Analyse the protein profile from the desired source, by separating and characterizing based on the molecular weight of the protein.

I Equipments Minigel/maxigel apparatus, power supply (capacity 200V, 500mA), boiling water bath, microcentrifuge and rocking/ rotatory shaker.

INTRODUCTION

1 Stock solutions 1 2M Tris-Hcl (pH 8.8) : 100ml. 2 1M Tris-Hcl (pH 6.8): 100ml. 3 10 %(w/v) SDS (store at room temperature). 4 50% (v/v) glycerol: 100ml. 5 1 %(w/v) bromophenol blue-10ml. (solution filtered after preparation to remove aggregated dye). 6 N, N, N’, N’-tetramethylene-ethylenediamine (TEMED). 7 2-mercaptoethanol or, dithiothreitol. 8 Glycine.

Sodium dodecyl sulphate polyacrylamide gel electrophoresis (SDS-PAGE) is used in a wide variety of applications for characterizing proteins. Few examples include analysis of antigen (protein) fractions from different sources for immunological and molecular biological studies, taxonomical applications by analysis of whole cell /cell envelope protein of the organism, and so on. Other types of electrophoresis in gels are listed in the table 51-1. Advantages and disadvantages of PAGE are mentioned in the box 51-1.

PRINCIPLE Electrophoresis is the migration of charged molecules in solution in response to an electric field. Their rate of migration depends on the strength of the field, on the net charge, size and shape of the molecules and also on the ionic strength, viscosity and temperature of the medium in which the molecules are moving. Polyacrylamide acts as a support matrix for running the sample. Sodium dodecyl sulphate (SDS) is an anionic detergent which denatures proteins by “wrapping around” the polypeptide backbone - and SDS binds to proteins fairly specifically; hence SDS confers a negative charge to the polypeptide in proportion to its length. The disulphide bridges in proteins are reduced with 2-mercaptoethanol.

II Reagents and lab wares Micropipette or Hamilton syringes (50µl and 100µl).

2 Working solutions Acrylamide stock (30%) :100ml. Acrylamide : 29.2g. Bisacrylamide : 0.8g. Add distilled water to make 100ml and stir until completely dissolved. 3 Separating gel buffer (4x) 75ml 2M Tris-Hcl (pH 8.8) 4ml 10% SDS 21ml distilled water

: 100ml. : 1.5M. : 0.4%.

4 Stacking gel buffer (4x) 50ml 1M Tris-HCl (pH 6.8) 4ml 10% SDS 46ml distilled water

: 100ml(Table 51-2). : 0.5M. : 0.4%.

5 10% Ammonium persulfate (APS) : 5ml. 0.5g APS dissolved in 5ml distilled water.

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6 Electrophoresis / Running Buffer (1x) : 1000ml. 3g Tris : 25mM. 14.4g glycine : 192mM. 1g SDS : 0.1%. Distilled water to make 1000ml 7 Sample buffer 0.6ml 1M Tris-HCl (pH 6.8) 5ml 50% glycerol 2ml 10% SDS 0.5ml 2-mercaptoethanol 1ml 1% Bromophenol blue 0.9ml distilled water

: 10ml. : 60mM. : 25%. : 2%. : 14.4mM. : 0.1%.

8 Staining solution 1.0g Coomassie blue R-250. 450ml methanol. 450ml distilled w.ater 100ml glacial acetic acid.

: 1000ml.

9 Destaining solution 100ml methanol. 100ml glacial acetic acid. 800ml distilled water.

: 1000ml.

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11 Connect the electrodes to the power pack and make prerun at a constant current of 20mA for 10-15minutes. 12 Switch off power supply and load the wells with your protein samples (10-50µl) diluted in sample buffer along with a molecular weight marker (5-10µl) using a micropipette/ syringe. 13 Run the electrophoresis at a constant current of 20mA till the dye front reaches the separating gel and then increases it to 25mA.Stop the run when the dye front reaches the bottom of the gel. 14 Now remove the side spacers from the glass plates and carefully scoop the gel using a spatula. The gel may be stained and destained for visualization or, used as such in blotting experiments without staining. 15 The gel may be stained using the Coomassie staining solution for 1-2hrs and then destained using the destaining solution for overnight in a rocking shaker. An alternate silver staining procedure too can be followed who is sensitive but a little expensive.

QUALITY CONTROL Known protein ladder is loaded in each run.

III Specimen Protein suspension to be separated.

OBSERVATIONS PROCEDURE 1 Mix protein sample (20µl) with sample buffer (5µl) in an Eppendorf tube and heat it at 75-100°C for 2- 10 minutes depending on the nature of the sample. 2 Assemble the clean glass plates by placing the spacerstwo on either sides, or one along the bottom edge and fix the whole assembly tightly with clamps or gel casting stand. 3 Seal the glass plate assembly with 2% agar or, melted wax on all three sides leaving the topside so that the assembly is leak proof. 4 Prepare and cast the separating gel using a small beaker. 5 Mix the components of the separating gel mixture without TEMED and de aerate. 6 Add TEMED finally to the mixture and pour immediately between glass plates upto 2cm below the notch and allow the gel to polymerize for 30-60 minutes at room temperature. 7 Overlay the acrylamide with n-butanol, which helps to keep the gel surface flat. After polymerization remove the butanol and rinse with water. 8 Prepare the stacking gel and cast over the separating gel as mentioned above and insert the comb. 9 Allow to polymerize for 15-30minutes. Then remove the comb carefully and rinse the wells with distilled water. 10 Now remove the casted gel plate from the gel casting stand/ clamps and detach the bottom spacer. Place the gel assembly into electrophoresis chamber with the notched plates facing inside and fill the upper and lower tanks with running buffer. Note: Remove air bubbles in the wells if any, using a syringe.

Observe the control protein bands. Observe and analyze the test protein bands.

RESULTS AND INTERPRETATION 1 The protein bands separated according to the molecular weight and charge are visualized directly. 2 Check out for the desired bands by comparing with a standard marker. 3 Interpret the desired band based on experimental and literature analysis by either visually or using a gel documentation system with appropriate software. 4 The bands can also be transferred by blotting and confirmed by coupling with appropriate antisera containing complimentary antibodies for the antigen (electro-immuno transfer blot).

Table51-1 List of other types of electrophoresis in gels Electroimmunodiffusion. Counterimmunoelectrophoresis. Rocket electrophoresis. Southern blotting. Northern blotting.

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Table 51-2 Calculation for X % separating or, stacking gel The gel concentration percentage, i.e.: whether 7.5, 10, 12.5 or 15 %, depends on ones experimental need and the desired quantity of stock solutions to cast the gel are calculated using the following formula. Acrylamide stock x /3 ml Stacking / separating gel buffer 2.5ml Distilled water (7.5-x /3) ml 10% APS 50µl TEMED 5µl (10µl if x < 8%) Total volume 10 ml 10% Separation gel Acrylamide stock Distilled water 10% APS TEMED Total volume

5% Stacking gel 3.3 ml. 4.2ml. 50µl. 5µl. 10 ml.

Acrylamide stock Distilled water 10% APS TEMED Total volume

1.6 ml. 5.9ml. 50µl. 5µl. 10 ml.

Separating gel buffer

2.5ml Stacking gel buffer

2.5ml

BOX 51-1 ADVANTAGES AND DISADVANTAGES OF PAGE Advantages Efficient tool for characterizing proteins. Highly sensitive. Rapid method. Protein bands are stable and can be stained for preservation. The number of different antigens can be readily observed. Disadvantages Chemicals are neurotoxic. Very expensive. Trained professional and well equipped laboratory is required.

VIVA 1 What is the principle of electrophoresis? 2 What are the applications of PAGE in microbiology? Ans.SDS-PAGE can be used a) in a wide variety of applications for characterizing proteins, b) in the diagnosis of infectious diseases, c) in analysis of antigen (protein) fractions, from different sources for immunological and molecular biological studies, d) in purification of antigen (protein) fractions, and e) in taxonomical applications by analysis of whole cell /cell envelope protein of the organism and categorizing accordingly. 3 Write a brief note on electro immuno transfer blot. Ans. Electro-immuno transfer blot (EITB), also known as Western blot is an immunological technique, which is used to detect a protein immobilized on a matrix. Prior to immunoblot, the protein of interest from a complex mixture is separated based on their molecular weight by sodium dodecyl sulfate – polyacrylamide gel electrophoresis (SDS-PAGE). EITB combines the resolution of gel electrophoresis with the specificity of immunochemical detection. The limit of detection by EITB is of the order 10 picogram with horse radish peroxidase labeling. Since the loading capacities of the gels are limited, an antigen cannot be detected when its concentration falls below 1 ng/sample. The EITB technique involves six steps: 1 Preparation of antigen sample. 2 Resolution of the sample by gel electrophoresis. 3 Transfer of the separated polypeptides to membrane support. 4 Blocking of nonspecific binding sites on the membrane. 5 Addition of antibody. 6 Detection. 4 What is the action of sodium dodecyl sulphate on proteins?

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KEY FACTS 1 2 3 4

Care must be taken for spillage or direct exposure to reagent while casting a gel. Avoid introducing air bubbles in to the gel. Always freshly prepared buffers and reagents should be used. Observe for complete circuit of electricity (use sufficient levels of buffer) throughout the procedure.

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Systemic Bacteriology. Volume 2.. Arnold publishers. pp. 2002, pp 1501. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. pp 94-96. 3 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. pp. 921.

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LESSON

52

Isolation of Antibiotic Resistant Mutant

LEARNING OBJECTIVES After completing this practical you will be able to:

method, colonies appearing in a region of high streptomycin concentration are indicative of streptomycin-resistant mutants. Different types of mutations and their role in microbial infections are summarized in the table 52-1.

1 Isolate the antibiotic resistant mutants from a prototrophic bacterial population.

REQUIREMENTS INTRODUCTION Mutation is any change in genetic information relative to a reference “wild-type” genome, including changes that affect expression of genes without altering their coding sequences and changes that do not cause any detectable phenotypic difference (silent mutations). In a complex organism, mutation can occur at many different structural levels and can be classified in many different ways. Mutations are mainly classified into two types: Point mutations and large-scale mutations (Box 52-1).Mechanisms of mutations are many (Box 52-2).

The importance of mutation Genes are stable repositories of the information needed for synthesis of all of the RNA and proteins in a living organism. Survival and stability of each species is dependent on faithful replication of genetic information for use by each new generation. However, a low level of mutational change is highly desirable. Over an extended period of time, mutational changes provide the ability for species to adapt to changing conditions and challenges. Hence, mutational changes serve as the raw material for selective survival and the evolution of more advanced and efficient species, as well as the development of biological diversity (Box 52-3).

PRINCIPLE After inoculation of the prototrophic test culture in to trypticase soy agar medium containing streptomycin by spread plate

I Equipments Micro centrifuge. II Reagents and lab wares Eppendorf tubes, micropipettes, two 10 ml trypticase soy agar in 20 ml test tubes and streptomycin solution (10 mg per 100 ml distilled water). III Specimen Pure growth of Escherichia coli from solid media preferably from non-blood agar plates (Examples: nutrient agar, MullerHinton agar), to be tested for antibiotic resistance is used as specimen.

PROCEDURE 1 Melt two trypticase soy agar tubes in a water bath and cool to 45°C. 2 Place a glass rod under one end of a sterile Petri dish, pour the molten agar until it cover the entire bottom surface and allow to solidify in the same position. 3 Add 0.1 ml of the streptomycin solution to a second tube of molten trypticase soy agar tube using a sterile pipette and mix gently. 4 Place the dish in horizontal position and pour the molten agar medium containing streptomycin medium until it fully covers the gradient agar layer and allow to solidify. 5 Add 0.2 ml of E. coli test culture with a sterile pipette and spread the culture with a sterile bent rod on entire agar surface.

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6 Incubate the plate in an inverted position for 48 hours at 37°C. 7 Select one or two isolated colonies from the middle of the streptomycin concentration gradient with a sterile loop and streak toward the high concentration end of the plate. 8 Incubate the plate for 24-48 hours at 37°C.

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Table 52-1 Different types of mutations and their role in microbial infections Type of mutation

Role in microbial infections

Point mutations Large scale mutations

Avian sarcoma/leukemia virus HIV virus Leishmania donovani Neisseria gonorrhoea MRSA Gyrase risistance gene in Salmonella Typhi

The parent strains (streptomycin sensitive strains) are included as controls.

RESULTS AND INTERPRETATION OBSERVATIONS Observe the colonies after first and second incubation steps.

1 Presence of colonies after first incubation indicates that the strain is resistant to streptomycin. 2 Sensitive strains do not produce colonies after first incubation. 3 Presence of colonies after second incubation denotes the resistant strains.

BOX 52-1 POINT MUTATIONS AND LARGER SCALE MUTATIONS In classical genetics, a point mutation was originally defined as a change in an inherited trait that was not accompanied by any chromosomal change that could be seen with a light microscope. Point mutation can result in missense (amino acid substitution), nonsense (insertion of a stop codon), or frameshift (either positive or negative). Missense mutations Most base pair substitutions change the amino acid specified by the codon in which they occur. Such mutations are described as missense mutations because they cause an amino acid substitution in the coded protein. Depending on the nature of the amino acid substitution and its location within the protein, missense mutations may have a variety of effects, ranging from complete loss of biological activity to reduced activity or temperature-sensitive activity or no functional effect at all. Nonsense mutations Base pair substitutions that generate an in-frame stop codon within a previously functional protein coding sequence cause premature termination of translation of the protein in question and are referred to as nonsense mutations. Silent mutations In some cases, base pair substitutions generate a different codon for the same amino acid, with no biological effect whatsoever. This is most likely to happen in the third position (wobble base) of redundant codons for the same amino acid. Such changes are considered to be mutations because they alter the genetic code. However, because they have no phenotypic effect, even at the level of protein amino acid sequence, they are called silent mutations. Frameshift mutations Addition or deletion of a single base pair in the middle of a coding sequence will result in out-of-frame translation of all of the downstream codons, and thus result in a completely different amino acid sequence, which is often prematurely truncated by stop codons (UAG, UAA, UGA) generated by reading the coding sequence out-of-frame. Such mutations, which are a special subclass of point mutations, are referred to as frameshift mutations. Deletion of a single base pair results in moving ahead one base in all of the downstream codons, and is often referred to as a positive frameshift. Addition of one base pair (or loss of two base pairs) shifts the reading frame behind by one base, and is often referred to as a negative frameshift. Large scale mutations Mutations that involve larger changes in chromosomes, including deletions, duplications, inversions, translocations from one chromosome to another, and extra or missing chromosomes are large-scale mutations.

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BOX 52-2 MECHANISMS OF MUTATION Tautomerization Spontaneous mutations that involve base pair substitutions are caused primarily by configurational changes within the individual bases that result in mispairing. These changes, which are called tautomeric shifts, involve momentary expression of rare alternative molecular configurations that exist in equilibrium with the more common forms. Specifically, proton shifts can convert the amino groups in adenine and cytosine to imino groups, and the keto groups in guanine and thymine to enol groups. Transitions A tautomeric shift in any of the four DNA bases can lead to mispairing of A to C or G to T. The tautomeric state can occur either in the template base or the incoming base. During the next round of DNA synthesis, the mispaired base will pair with its normal partner, resulting in a transition, in which an AT base pair replaces a GC or a GC replaces an AT, with no change in the purine: pyrimidine polarity of the base pair. Transitions are the most common type of mutation resulting from spontaneous mispairing due to tautomerization. Transversions To achieve a transversion, in which the positions of purine and pyrimidine are reversed in the DNA double helix, two events are thought to be involved, tautomerization of one of the bases and rotation of the other to yield a purine: purine pairing. The frequency of spontaneous transversions, which is lower than that of transitions, appears to be consistent with this interpretation. A second possible mechanism for transversions is the formation of an apurinic site, which can result in replacement of the original purine with any of the four bases.

BOX 52-3 THE IMPORTANCE OF MUTATION Genes are stable repositories of the information needed for synthesis of all of the RNA and proteins in a living organism. Survival and stability of each species is dependent on faithful replication of genetic information for use by each new generation. However, a low level of mutational change is highly desirable. Over an extended period of time, mutational changes provide the ability for species to adapt to changing conditions and challenges, and thus serve as the raw material for selective survival and the evolution of more advanced and efficient species, as well as the development of biological diversity.

KEY FACTS 1 Properly mix streptomycin solution to molten trypticase soy agar. 2 Sufficiently cool the molten agar before adding antibiotic solution. 3 Presence of colonies after second incubation denotes the resistant strains.

VIVA 1 Define and classify mutations. 2 Write a short note on point mutations. 3 What are the mechanisms of mutations? Ans. Tautomerization, transitions, and transversions. 4 Explain the mechanism of isolation of antibiotic resistant mutant strains. 5 What are different types of mutations and their role in microbial infections? FURTHER READING 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Systemic Bacteriology. Volume 2. Arnold publishers. pp. 2002, pp 1501. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. pp 94-96. 3 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. pp. 921.

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LESSON

53

Bacterial Conjugation

LEARNING OBJECTIVES

Differences between F+ strain and Hfr strain are listed in the Table 53-1.

After completing this practical you will be able to: 1 Demonstrate genetic recombination in bacteria by conjugation.

INTRODUCTION Bacterial conjugation can be viewed as a primitive form of sex, in which a cell from a donor strain injects DNA into a recipient cell, where it can undergo recombination and become part of the recipient’s genome. Donor strains contain an additional genetic element, called the fertility factor (F), usually in the form of a plasmid. Cells that are receptive to conjugation lack the F factor and are sometimes designated F –. During conjugation of an F + cell with an F –, the frequency of transformation for any genes other than the F factor is very low. However, if it does happen, it is due to random chromosomal integration of the F factor.

Hfr strains In certain F+ strains, the F factor has become integrated into the bacterial chromosome. When this happens, transfer of chromosomal DNA from the donor to the recipient F– cell begins adjacent to the integrated F factor and can progress around the entire bacterial chromosome if the process is not interrupted. Bacterial strains with integrated F factors are referred to as Hfr strains (“high frequency of recombination”).

F’ factors and sexduction Sometimes an integrated F+ factor from an Hfr strain will escape from the bacterial chromosome carrying a few chromosomal genes with it in its circular plasmid DNA. Such a plasmid is called an F-prime (F’) factor. The bacterial genes carried on the F’ plasmid are easily transmitted into F– cells by conjugation.

PRINCIPLE After inoculation and incubation of F– and Hfr strains together in a minimal growth medium containing thiamine and streptomycin, threonine and leucine, recombinant strains of the bacteria only grow. Thiamine acts as a nutrient supplement to thiamine negative recombinant cells. Streptomycin inhibits the growth of parental streptomycin sensitive Hfr strains. Parental F– strains cannot grow due to the lack of threonine and leucine. Mechanisms of DNA transfer are highlighted in the Box 53-1.

REQUIREMENTS I Equipments Incubator. II Reagents and glass wares Bend glass rod, sterile pipettes, and test tubes, 24 hour nutrient broth cultures of streptomycin resistant F– Escherichia coli strain which require threonine, leucine and thiamine, and streptomycin sensitive Hfr E. coli strain, nutrient agar plates and 95% ethanol. III Specimen F– E. coli culture.

PROCEDURE 1 Take nutrient agar medium. 2 Add 1 ml of F– E. coli culture and 0.3 ml of Hfr E. coli cultures into a sterile test tube. 3 Incubate for 30 min at 37°C. 4 Vigorously agitate the mixed culture to terminate the genetic transfer.

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Bacterial Conjugation

5 Add 0.1 ml of the mixed culture to a minimal growth medium containing streptomycin and thiamin and spread over the entire surface agar surface. 6 Incubate the plates for 48 hours at 37°C.

Observe the culture plates for colonies.

QUALITY CONTROL

RESULTS AND INTERPRETATION

Prepare control plates of parental Hfr and F– and aseptically add 0.1 ml of each strain to its agar plate and spread with a bent gloss rod.

Presence of colonies in the test medium indicates the transfer of genetic material.

OBSERVATIONS

BOX 53-1 MECHANISMS OF DNA TRANSFER The DNA transfer that occurs in conjugation begins as a single strand break in the donor chromosome (or plasmid), with only one strand transferred through the F– pilus to the recipient cell. The single strand left behind in the donor and the one that is transferred to the recipient are both converted into double strands, restoring the donor chromosome and generating double stranded donor DNA in the recipient. As a result of receiving new DNA from the donor, the recipient is temporarily partially diploid. Recombination can then integrate parts of the transferred DNA into the recipient genome. Any DNA that is not integrated is soon destroyed. Transformation Transformation refers to the ability of extracellular DNA to enter a bacterial cell and recombine with the bacterial genome, thereby giving the bacterial cell new genetic properties. Transduction Transduction refers to a genetic exchange in which bacteriophages carry bacterial genes from one bacteria cell to another. There are two classes of transduction, specialized and generalized. In specialized transduction, a lysogenic phage undergoes recombination with the host genome and later when it is excised to become an independent phage genome, it carries one or more host genes with it, which can be transduced into a new host cell and recombined into the genome of that cell. In generalized transduction, fragments of host DNA are mistakenly packaged into a bacteriophage in place of its own DNA.

Table53-1 Differences between F+ strain and Hfr strain F+ strain

Hfr strain

1. F factor usually present in the form of a plasmid. 2. Low frequency of recombination. 3. Only F factor transfer occurs in conjugation.

1. F factor has become integrated into the bacterial chromosome. 2. High frequency of recombination. 3. Transfer of chromosomal DNA from the donor to the recipient F– cell begins adjacent to the integrated F factor and can progress around the entire bacterial chromosome if the process is not interrupted.

KEY FACTS 1 Young cultures of bacterial strains must be used. 2 Mix the culture properly to terminate the genetic transfer. 3 Avoid contamination.

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VIVA 1 2 3 4

Define bacterial recombination. What are the methods of transfer of genetic material in bacteria? How do you demonstrate bacterial conjugation? What are the applications of bacterial recombination? Ans: Applications of bacterial recombination include: a Production of recombinant antigens for diagnosis. b Production of recombinant vaccine proteins. c Gene therapy. d Transfer of antibiotic resistance, and e Production of recombinant strains of bacteria with required characteristics. 5. Differentiate between F+ and Hfr strains. 6. Discuss the role of conjugation in acquiring drug resistance. Ans: Plasmids carry only a few genes, often for antibiotic resistance. Plasmids that carry appropriate genes are capable of making bacteria resistant to antibiotics. The plasmid is infectious and can transfer itself by conjugation. The plasmid (F factor) is present as an independent double stranded ring of DNA distinct from chromosome. During conjugation there is a break in one of the two strands of the plasmid. This strand passes in to the recipient cell. Donor cell retains one strand. Donor and recipient cells synthesize complimentary strands to form intact duplexes and recipient cell becomes a potential donor cell. The F1 factor is formed by the excision of the F factor from the chromosome along with a segment of the chromosome. The release of F factor occurs by breakage and reunion at a point different from the point of plasmid insertion. This results in an interchange of plasmid and chromosomal DNA segments.

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Systemic Bacteriology. Volume 2. Arnold publishers. pp. 2002, pp 1501. 2 Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 1995. pp 94-96. 3 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. pp. 921.

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UNIT

VIII Bacteriology

Lesson 54 Normal Microbial Flora of the Mouth Lesson 55 Normal Microbial Flora of the Throat Lesson 56 Normal Microbial Flora of the Skin Lesson 57 Identification of Staphylococcus aureus Lesson 58 Identification of Streptococcus pneumoniae Lesson 59 Identification of b-haemolytic Streptococci Lesson 60 Identification of Corynebacterium diphtheriae Lesson 61 Identification of Lactose Fermenting Enterobacteriaceae Lesson 62 Identification of Vibrio cholerae Lesson 63 Identification of Pseudomonas aeruginosa

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LESSON

54

Normal Microbial Flora of the Mouth

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate various microorganisms which are present as part of the normal flora in the mouth.

INTRODUCTION Normal flora usually denotes to microorganisms, which inhabit the skin and mucous membrane of the normal human body. Within a few hours of birth, normal oral and nasopharyngeal flora appear. The normal microbial flora offers many benefits to the human (Box 54-1). There are many microorganisms which are present as part of the normal flora of the mouth and teeth. These are Streptococcus viridans, Streptococcus mutans, nonpathogenic Neisseria, Staphylococcus, Fusiform species, anaerobic spirochaetes, etc. In infant at the time of birth, the mouth contains usually the organisms that are present in mother’s vagina (e.g., micrococci, streptococci, coliform bacilli, Döderlein’s bacilli, etc.), as they acquire these organisms during their birth through the vaginal canal. These organisms however diminish in number during first week after birth and are replaced by the normal bacterial flora present in the mouth of the mother. Some of these organisms can cause opportunistic infections. Many of these bacteria can be demonstrated in the saliva.

II Reagents and lab wares Petri dishes, sample collection bottle, glass slides, pencil, inoculation loop, and burner, Gram stain reagents (methyl violet, iodine, 95% alcohol and dilute carbol fuchsin), filter paper containing 1% tetramethyl paraphenylene diamine dihydrochloride, two blood agar and chocolate agar plates. III Specimen Saliva.

PROCEDURE 1 Collect the saliva into a sterile sample collection bottle. 2 Inoculate the saliva onto blood agar and chocolate agar plates. 3 Incubate the chocolate agar plate in CO2 incubator for 48 hours at 37°C. 4 Incubate one blood agar plate aerobically for 48 hours at 37°C.

QUALITY CONTROL 1 Test all agar plates for sterility before inoculation. 2 Incubate uninoculated blood agar and chocolate agar plates along with the inoculated ones.

PRINCIPLE

OBSERVATIONS

Saliva is collected from the mouth and inoculated onto the surface of the agar plates. The colonies grown on these plates are studied further for their identification.

1 Look for the presence of haemolysis on blood agar plate by viewing the plate under transmitted light. 2 Observe all the colonies on both plates. 3 Record your observations. 4 Perform Gram stain. Observe and record morphology of the bacteria.

REQUIREMENTS I Equipments Microscope and CO2 incubator.

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RESULTS AND INTERPRETATION 1 On blood agar streptococci produce pinpoint a-hemolytic colonies where as staphylococci produce pinhead colonies with b - haemolysis. 2 On Gram stain, streptococci are identified based on the arrangement, they are Gram positive cocci arranged in chains. Staphylococci are arranged in clusters. 3 On chocolate agar, colonies grown are tested for the presence of the enzyme oxidase (refer chapter 21). Neisseria are oxidase positive. They turn the oxidase paper into deep purple in color when they are streaked on the surface of the filter paper.

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BOX 54-1 BENEFICIAL EFFECTS OF NORMAL FLORA 1 They suppress the colonization of the body by pathogens. 2 Colicins produced by some bacteria lyse some pathogenic bacteria. 3 Antibodies produced against some commensals show cross reaction with some pathogens, thereby enhancing immune status of the host. 4 Endotoxins produced by some bacteria facilitate complement mediated defence system of the humans.

KEY FACTS 1 There are many microorganisms which are present as part of the normal flora in the mouth and teeth. 2 Saliva sample should be collected in a sterile bottle. 3 Saliva sample should be processed immediately after collection.

VIVA 1 What are the microorganisms present in the mouth as normal flora?

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002.

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LESSON

55

Normal Microbial Flora of the Throat

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate various microorganisms which are present as part of the normal flora in the throat and respiratory tract.

INTRODUCTION Many microorganisms are present in the throat and upper respiratory tract. These are a-hemolytic streptococci, Streptococcus pneumoniae, Staphylococcus, Branhamella, Neisseria, Haemophilus, diphtheroids and spirochaetes. a- haemolytic streptococci are present in the upper respiratory tract within 12 hours of the birth. They continue to remain as part of important microbial flora through out life. The microbial flora present in the pharynx and trachea are similar to that present in the mouth.

III Specimen Throat swab.

PROCEDURE 1 Place a tongue depressor on the extended tongue and collect the specimen by using a sterile cotton swab, by rotating the cotton swab gently over pharyngeal tonsils. 2 Emulsify the throat swab in sterile saline and mix to form a uniform suspension. 3 Inoculate the specimen onto blood agar, chocolate agar and potassium tellurite agar plates. 4 Incubate the chocolate agar plate in CO2 incubator for 48 hours at 37°C. 5 Incubate one blood agar plate and potassium tellurite agar plates aerobically for 48 hours at 37°C.

OBSERVATIONS PRINCIPLE Specimen taken from the throat is inoculated onto the surface of the media plates. The colonies grown on these plates are studied further for their identification.

REQUIREMENTS I Equipments Microscope and CO2 incubator. II Reagents and lab wares Bunsen burner, tongue depressor, sterile cotton swabs, glass slides and pencil, Gram staining reagents (methyl violet, Iodine, 95% alcohol and dilute control fuchsin), filter paper containing 1% tetramethyl paraphenyline diamine dihydrochloride, two blood agar plates, one potassium tellurite agar plate, and one 5ml sterile saline tube.

1 Look for the presence of hemolysis on blood agar plate by viewing the plate under transmitted light. 2 Perform oxidase test for the colonies grown on chocolate agar. 3 Look for the presence of black colored colonies on potassium tellurite agar. 4 Record your observations. 5 Perform Gram stain. Observe and record morphology of the bacteria.

RESULTS AND INTERPRETATIONS 1 On blood agar, streptococci produce pinpoint a-hemolytic colonies where as staphylococci produce pinhead colonies with b-hemolysis. 2 On Gram stain, streptococci are identified based on the arrangement, they are Gram positive cocci arranged in chains. Staphylococci are arranged in clusters.

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3 On chocolate agar, colonies grown are tested for the presence of the enzyme oxidase (refer chapter 21). Neisseria are oxidase positive. They turn the oxidase paper into deep purple in color when they are streaked on the surface of the filter

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paper. Oxidase negative bacteria do not produce color when the colonies are streaked on oxidase paper. 4 Diphtheroids when grown on potassium tellurite agar produce pinpoint black colored colonies.

KEY FACTS 1 There are many microorganisms which are present as part of the normal flora in the throat and upper respiratory tract. 2 Throat swab should be collected in a sterile bottle. 3 Throat swab should be processed immediately after collection.

VIVA 1 In which respect, the normal microbial flora of human is helpful? Ans. The normal microbial flora of human can reduce the infections by competing with pathogens for nutrients and binding receptors. Some organisms also form symbiotic association with the host.

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002.

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LESSON

56

Normal Microbial Flora of the Skin

LEARNING OBJECTIVES

dextrose agar plate, lactophenol cotton blue and reagents for Gram stain.

After completing this practical you will be able to: 1 Demonstrate various microorganisms which are present as part of the normal flora on the skin.

III Specimen Skin swab.

INTRODUCTION

PROCEDURE

Skin as part of its normal flora, normally harbours many bacteria. It contains 102 to 104 organisms per cm2. Staphylococcus epidermidis and diphtheroids are predominant bacteria. The microbial flora present in the pharynx and trachea are similar to that present in the mouth. Streptococcus viridans, micrococci, peptostreptococci, propionibacterium, b-haemolytic streptococci, enterococci, diphtheroids, Candida species, Malasezia furfur , etc. are the other microorganisms.

1 Collect the swab from surface of the skin by rubbing the cotton swab after moistening the swab in sterile saline. 2 Place the swab in a tube containing sterile saline and mix it. 3 Inoculate a loopful suspension on one plate each of blood agar, mannitol salt agar and Sabouraud’s dextrose agar 4 Incubate the Sabouraud’s agar plate for 48 hours at 25°C and the remaining plates for 48 hours at 37°C.

OBSERVATIONS PRINCIPLE

REQUIREMENTS

1 Look for the presence of hemolysis on blood agar by viewing the plates under transmitted light. 2 Look for the presence of mold like or moist growth on Sabouraud’s agar plate. 3 Look for the presence of yellow color of the medium surrounding colonies grown on mannitol salt agar medium. Yellow color colony is suggestive of S. aureus.

I Equipments Microscope and incubator.

RESULTS AND INTERPRETATION

II Reagents Bunsen burner, sterile cotton swabs, glass slides and pencil, blood agar plate, mannitol salt agar plate, Sabouraud’s

1 Perform Gram stain of the colonies. 2 Perform LPCB stained smears of colonies grown on Sabouraud’s dextrose agar. 3 Record your observations.

Swab is collected from surface of the skin and is inoculated on agar plates and incubated. The colonies are studied further for their identification.

VIVA 1 What is the role of resident flora of skin?

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KEY FACTS 1 Skin as part of its normal flora, normally harbours many bacteria. 2 The normal resident flora of skin can reduce the infections by competing with pathogens for binding receptors.

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002.

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LESSON

57

Identification of Staphylococcus aureus

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know about the clinical importance of staphylococci. 2 Know the procedures used to identify Staphylococcus aureus and differentiate it from other staphylococcal species.

INTRODUCTION Staphylococcus species are Gram positive spherical shaped bacteria measuring 0.8 µm to 1 µm in size arranged in clusters. They are catalase positive, non-spore forming bacteria Staphylococci are part of the indigenous flora of skin surfaces, mucous membranes and upper respiratory tract. They usually cause suppurative lesions such as pustule, furuncle or carbuncle and can also involve muscles and bones. They are less commonly associated with systemic infection. However they produce a variety of exotoxins causing various toxin-mediated diseases such as food poisoning, toxic shock syndrome and staphylococcal scalded skin syndrome. Laboratory tests for differentiation of staphylococcal species are listed in the table 57-1.

SPECIMENS Clinical specimens like pus swab, and wound swab. In this chapter the following tests will be described which are employed routinely for identification of S. aureus.

TESTS FOR THE IDENTIFICATION OF STAPHYLOCOCCUS AUREUS 1 Direct examination Gram stain Gram stain of smears of clinical specimens is carried out to

demonstrate the presence of Gram positive cocci arranged in clusters along with pus cells (Fig. 57-1).

2 Culture Clinical specimens are inoculated onto nutrient agar, blood agar and MacConkey agar plates and kept for aerobic incubation at 37°C. S. aureus produces opaque, circular colonies with butyrous consistency. Golden yellow pigment is demonstrated on nutrient agar (Fig. 57-2). On blood agar, they produce bhaemolytic golden yellow or white colonies (Fig. 57-3). On MacConkey agar they produce minute pink colored colonies.

3 Coagulase test Coagulase test is most important test used for identification of S. aureus (refer chapter 22).

4 Deoxyribonuclease test Coagulase positive S. aureus also produce the enzyme deoxyribonuclease enzyme. Detection of the presence of the enzyme, deoxyribonuclease is used to reconfirm the identification of S. aureus. The test organism is streaked onto the DNA agar medium (0.2% DNA). Then the DNA plate is incubated overnight at 37°C. After incubation, 3.6% of hydrochloric acid (1N HCl) is added to the medium to determine the DNase activity. DNase positive S. aureus hydrolyze the DNA resulting in production of halo around the growth. Absence of halo around the growth indicates DNase-negative S. aureus which are also coagulase negative.

5 Mannitol salt agar This is a selective medium used to isolate staphylococci from clinical specimens. It contains nutrient agar with 1% mannitol, and 7.5% sodium chloride. Phenol red is used as an indicator. The medium is used to test mannitol fermenting ability of

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S. aureus. It also tests ability of S. aureus to tolerate the high salt concentration in the medium and grow readily. S. aureus produces yellow colonies on the medium while other staphylococcal species do not. The yellow colour is seen due to the fermentation of mannitol with production of acid by S. aureus, thereby reducing pH of the medium to the acidic. In acidic pH the indicator, phenol red, produces yellow colour.

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The plate is then incubated overnight at 37°C. The presence of zone of inhibition (17 mm) around the disc indicates novobiocinsensitive S. aureus and S. epidermidis and no zone of inhibition around the disc indicates novobiocin-resistant S. saprophyticus.

6 Novobiocin sensitivity Novobiocin sensitivity testing is used to detect sensitivity or resistance of S. aureus to the antibiotic, novobiocin. This test is used to differentiate S. aureus and S. epidermidis from S. saprophyticus. Both S. aureus and S. epidermidis are novobiocinsensitive where as S. saprophyticus is novobiocin-resistant. In this test, staphylococci strain to be tested is inoculated onto the surface of a Mueller Hinton agar plate, followed by application of a 5µg novobiocin disc on the surface of the agar.

FIGURE 57-1 Pus smear showing Gram positive cocci in clusters, x 1000.

Table 57-1 Laboratory tests for differentiation of staphylococcal species Test

S. aureus

S. epidermidis

S. saprophyticus

Growth on Mannitol salt agar Colonial pigmentation Coagulase test DNAase test Hemolysis on blood agar Novobiocin sensitivity

+ Golden yellow + + Beta Sensitive

White Sensitive

White Resistant

FIGURE 57-2 Staphylococcus aureus colonies on nutrient agar.

FIGURE 57-3 Staphylococcus aureus colonies on blood agar.

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Identification of Staphylococcus aureus

KEY FACTS 1 Yellow color pigmentation can also be produced by micrococci which can be distinguished from staphylococci by various biochemical tests such as modified oxidase and sensitivity for novobiocin, bacitracin and furazolidone. 2 Some coagulase negative staphylococci can produce beta hemolysis on blood agar similar to coagulase positive staphylococci, hence should not be confused with that of S. aureus. 3 Some species of staphylococci produce only bound coagulase (clumping factor) such as S.hydenensis and S.schleifer

VIVA 1 What are the tests to identify S. aureus? 2 List novobiocin-sensitive staphylococci.

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002.

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LESSON

58

Identification of Streptococcus pneumoniae

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know about the clinical importance of pneumococci. 2 Know the procedures used to identify Streptococcus pneumoniae and differentiate it from other alpha hemolytic streptococci.

INTRODUCTION S. pneumoniae are Gram positive cocci arranged in pairs and individual coccus is lanceolate shaped. They are encapsulated and are an important cause of lobar pneumonia. They also cause meningitis and other infections such as otitis media, conjunctivitis, peritonitis, sinusitis and suppurative arthritis.

SPECIMENS Clinical specimens such as sputum, pleural exudate, CSF, blood, etc. S. pneumoniae produce a haemolysis on blood agar. S. pneumoniae can be differentiated from other alpha hemolytic streptococci by many tests (Table 58-1). In this chapter the following tests will be described which are employed routinely for identification of S.pneumoniae.

TESTS FOR THE IDENTIFICATION OF STREPTOCOCCUS PNEUMONIAE 1 Direct examination Gram stain Gram stain of smears from clinical specimens is carried out to demonstrate the presence of Gram positive cocci. These are

small, slightly elongated cocci arranged in pairs (diplococci) with the broad ends in apposition. Each coccus has one end pointed and other end broad or rounded (lanceolate shaped). Gram stain of the CSF specimen from a case of meningitis shows both intracellular as well as extra cellular cocci.

2 Culture The specimens are inoculated onto blood agar and kept for aerobic incubation at 37°C for 24 hours in the presence of C02. S.pneumoniae produces small, shiny dome-shaped and translucent colonies which are surrounded by alpha hemolysis (Fig. 58-1). On prolonged incubation, these colonies may show a depressed center with an elevated rim (Draughtsman’s colonies). This is due to the autolysis of the bacteria in old colony.

3 Bile solubility test S. pneumoniae undergo autolysis in the presence of surface active agents such as sodium deoxycholate whereas other alpha hemolytic streptococci are bile insoluble hence is not lysed by these bile salts. Label two brain heart infusion broth tubes, one as S. pneumoniae (positive control) and the other tube as S. mitis (negative control). Add loopful of test organisms in the broth to give a heavy suspension. Then add 0.5 ml of sodium deoxycholate solution into each tube. Incubate the tubes in a water bath at 37°C for one hour. Clearing of turbidity indicates positive test whereas persistence of turbidity indicates negative test. S. pneumoniae is bile solubility positive.

4 Optochin test S. pneumoniae are sensitive to Optochin (ethyl hydrocupreine hydrochloride) and produce a zone of inhibition measuring 15 mm and more. Other alpha hemolytic streptococci are resistant to Optochin which produce a zone of inhibition of less than 15 mm (Fig. 58-2).

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Identification of Streptococcus pneumoniae

A blood agar plate is divided into two halves by glass marking pencil. One half is labeled as S. pneumoniae while the other half is labeled as S. mitis. Using a sterile cotton swab, test strains are inoculated onto the surface of blood agar, followed by application of 0.05 units Optochin disc over the inoculated surface by using forceps. The plate is incubated aerobically at 37°C under 10% CO2 environment for 24 hours. Development of zone of inhibition of 15 mm and more shows the test to be positive. Zone of inhibition less than 15 mm shows the test to be negative.

5 Inulin fermentation

FIGURE 58-1 Streptococcus pneumoniae on blood agar.

FIGURE 58-2 Optochin sensitivity test.

S. pneumoniae is capable of fermenting the sugar, inulin whereas other alpha hemolytic streptococci do not ferment inulin. The test strain is inoculated into the inulin sugar medium (serum sugar) and is incubated over night at 37°C. Positive test is indicated by change of color of the media from red to yellow. Negative test is indicated by no color change, and medium continues to appear as red.

Table58-1 Differences between Streptococcus pneumoniae and Streptococcus mitis Tests 1. Bile solubility 2. Optochin sensitivity 3. Inulin fermentation 4. Quellung reaction 5. Mouse virulence

Streptococcus pneumoniae + + + + +

Streptococcus mitis -

KEY FACTS 1 Streptococcus pneumoniae are Gram positive cocci arranged in pairs or as short chains and individual coccus is lanceolate shaped. 2 Gram stain of the CSF from a case of meningitis shows both intracellular as well as extra cellular cocci. 3 Str. pneumoniae is bile solubility test positive. 4 Str. pneumoniae are sensitive to Optochin. 5 Str. pneumoniae is capable of fermenting the sugar, inulin whereas other alpha hemolytic streptococci do not ferment inulin.

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VIVA 1 How do you demonstrate capsules of S. pneumoniae? Ans. India ink preparation. 2 What is the mechanism of autolysis? Ans. Intracellular autolytic enzyme mediated. 3 What is the strength of the routinely used Optochin disc? 4 What are the rapid diagnostic methods for pneumococcal meningitis? Ans. Capsular antigen detection in the CSF by latex agglutination and CIEP.

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002.

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LESSON

59

Identification of b-haemolytic Streptococci

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know about clinical significance of b-haemolytic streptococci. 2 Perform important tests to identify group A streptococci, group B streptococci and group D streptococci.

streptococci cause neonatal infections such as septicemia and meningitis. Group D streptococci may cause urinary tract infections and wound infections.

SPECIMENS Throat swab. In this chapter the following tests will be described which are employed routinely for identification of S. pyogenes.

INTRODUCTION All members of b-haemolytic streptococci are Gram positive and fastidious. They are cocci arranged in chains. They appear as circular translucent pinpoint colonies on blood agar. Streptococci, both aerobic and anaerobic are classified on the basis of their haemolysis on blood agar. On blood agar three types of haemolysis are observed. These are: a) a hemolysis: It is an incomplete form of hemolysis where a green zone is produced around the colonies. b) b hemolysis: It is a complete form of hemolysis. This appears as a clear zone around the colonies. b-haemolytic streptococci are frequently associated with pathogenicity, and c) g haemolysis: It is described as absence of any hemolysis around the colony. g-haemolytic streptococci are avirulent. b-haemolytic streptococci produce a clear zone of haemolysis on the blood agar surrounding the colonies, within which erythrocytes are completely lysed. The lysis is caused by two haemolysins, streptolysin O and streptolysin S. b-haemolytic streptococci on basis of their carbohydrate C antigen are classified in to 20 groups (from A to V, excepting I and J). This classification is known as Lancefield classification. Group A streptococci, also known as Streptococcus pyogenes is the important species causing most human infections. S. pyogenes causes a wide variety of suppurative infections of the respiratory tract, skin, genital and other deep infections. It also causes non-suppurative infections such as rheumatic fever and acute glomerulonephritis. Group B

TESTS FOR IDENTIFICATION OF STREPTOCOCCUS PYOGENES 1 Direct examination Gram stain Gram stain of smears from clinical specimens like throat swab is carried out to demonstrate the presence of Gram positive cocci arranged in short chains (Fig. 59-1).

2 Culture The specimens are inoculated onto 5% sheep blood agar and kept for aerobic incubation at 37°C for 24 hours in the presence of C02. Str.pyogenes produces pinpoint colonies which are surrounded by a large zone of b-hemolysis (Fig. 59-2).

3 Bacitracin sensitivity test Bacitracin test is a frequently used test to identify and differentiate group A streptococci from non- Group A streptococci. Bacitracin disc when placed on the surface of blood agar plate streaked with test organism inhibits its growth and forms a zone of inhibition. The test organism is streaked on to the surface of the blood agar. Then a bacitracin having strength of 0.04 units is placed on the inoculated plate. The plate is incubated at 37°C for

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overnight. Positive test is indicated by a zone of inhibition around the bacitracin disc. The negative test is indicated by no zone of inhibition around the bacitracin disc. Most strains of S. pyogenes are bacitracin sensitive.

4 CAMP (Christie, Atkins and Munch-Peterson) test

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when reacts with iron salts in the medium causes brown to black discolouration of medium following incubation. Test strain is inoculated onto the surface of the bile aesculin agar and kept for aerobic incubation at 37°C for 24–48 hr. Positive test is indicated by change of the brown colour of the medium in to black. Negative test is indicated by no brown or black discoloration of medium.

CAMP (Christie, Atkins and Munch-Peterson) test is a test used to identify Group B streptococci. CAMP substance is a peptide produced by group B streptococci which acts synergistically with the b-hemolysis produced by some strains of Staphylococcus aureus enhancing the effect of haemolysis. The test is performed by inoculating S. aureus strain as a straight line on the surface of the sheep blood agar. Known positive control (Group B streptococci), negative control (group A streptococci) and test strains are also inoculated as a straight lines parallel to S. aureus streak line leaving 1 cm space. The plate is incubated aerobically at 37°C overnight in 5–10% CO2 environment. Positive test is indicated by an arrow head zone of hemolysis near the S. aureus growth. Negative test is indicated by the absence of arrow head zone of hemolysis (Fig. 59-3).

5 Bile aesculin test This test is carried out to identify Group D streptococci. The cocci hydrolyse aesculin into 6, 7 dihydroxy-coumarin which

FIGURE 59-2 Beta hemolytic streptococci colonies on blood agar.

FIGURE 59-1 Streptococci in short chains, x 1000.

FIGURE 59-3 CAMP Test.

VIVA 1 2 3 4

What are different types of haemolysis produced by streptococci? What is the principle of the bacitracin sensitivity test? What is the principle of the CAMP (Christie, Atkins and Munch-Peterson) test? What is the principle of the bile aesculin test?

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Identification of b-haemolytic streptococci

KEY FACTS 1 b-haemolytic streptococci produce a clear zone of haemolysis on the blood agar surrounding the colonies, within which erythrocytes are completely lysed. 2 Bacitracin test is a frequently used test to identify and differentiate group A streptococci from non- Group A streptococci. 3 Blood agar plate for CAMP test should be incubated in the environment of 5–10% CO2. 4 CAMP test should be done with parallel positive and negative control strains. 5 Bile aesculin test is carried out to identify Group D streptococci, heat resistant test and salt tolerance test.

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002

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LESSON

60

Identification of Corynebacterium diphtheriae

LEARNING OBJECTIVES After completing this practical you will be able to 1 Know about the clinical importance of Corynebacterium diphtheriae. 2 Know the procedures used to identify C. diphtheriae and differentiate it from other non pathogenic Corynebacterium species.

INTRODUCTION C. diphtheriae are thin, non-sporing, non-capsulated and nonmotile Gram positive bacilli which measure approximately 3 µm × 0.3 µm in size. These bacilli have typical club-shaped swellings at one or both ends of bacilli. They are arranged in angled pairs or in angular fashion which resemble Chinese letters. These bacilli have metachromatic granules towards the end of the logarithmic growth period. These granules are the storage depots of polymetaphosphate. C. diphtheriae causes natural infection only in man. They produce diphtheriae toxin that causes local tissue necrosis which results in the formation of grayish white membrane at the affected site. Commonest sites affected are tonsils, posterior pharyngeal wall, nasal passages, larynx and trachea. Though diphtheria is a disease of respiratory tract this can also affect skin, conjunctiva, ear, etc.

Albert’s stain Albert’s stain is done to detect the presence of metachromatic granules in C. diphtheriae. The specimen is collected from the surface of the pseudomembrane by a sterile cotton swab. The swab containing the specimen is rolled on the surface of the clean glass slide. The smear is heat fixed and Albert’s stain is carried out (refer chapter 9). The smear is examined under oil immersion (100 x) objective (Fig. 60-1).Presumptive identification of C. diphtheriae is made based on the presence of metachromatic granules by Albert’s stain

2 Culture Loeffler’s serum slope and potassium tellurite agar are the two media frequently used for culture of C. diphtheriae. Loeffler’s serum slope is inoculated with the throat swab and incubated at 37°C for 6 hours. C. diphtheriae produces minute smooth colonies after 6 hours of incubation (Fig. 60-2). Colonies grown on this medium are stained by Albert’s staining for demonstrating metachromatic granules. Similarly, the swab specimen is inoculated on the surface of Macleod’s potassium tellurite agar media (0.04%) and is incubated at 37°C for up to 48 hours. C. diphtheriae reduces potassium tellurite into metallic tellurium which results in production of black colored colonies on the medium (Fig. 60-3).

3 Biochemical tests SPECIMENS Throat swab. In this chapter the following tests will be described which are employed routinely for identification of C. diphtheriae.

TESTS FOR THE IDENTIFICATION OF CORYNEBACTERIUM DIPHTHERIAE 1 Direct examination

The following biochemical tests are carried out: a. Fermentation of serum sugars (glucose, maltose, sucrose, lactose, mannitol, and trehalose) for the detection of acid. The colonies grown on blood agar are inoculated on to serum sugars and gelatin, and incubated at 37°C for 24 hours. The colour change in the serum sugar media is looked for. Positive test is indicated by the change of media to yellow colour due to production of acid. Negative test is indicated by the persistence of red colour, because no acid is produced. C. diphtheriae ferments glucose, maltose and sucrose with

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production of acid only. They do not ferment lactose, mannitol, and trehalose. b. Other biochemical tests which are used primarily to differentiate C. diphtheriae from other Corynebacterium spp. include urea hydrolysis and gelatin hydrolysis. Positive urea hydrolysis test is indicated by change in colour of the medium into red. Negative urea hydrolysis test is indicated by no colour change. Positive gelatin hydrolysis test is indicated by free flowing of medium on inverting the gelatin tube. No free flowing on inversion of tube indicates negative test. C. diphtheriae do not hydrolyse urea and gelatin whereas non-pathogenic Corynebacterium species such as C. ulcerans hydrolyses urea.

FIGURE 60-2 Loeffler’s serum slope.

FIGURE 60-1 Albert’s stain showing volutin granules, x 1000. FIGURE 60-3 Black coloured colonies on potassium tellurite agar.

KEY FACTS 1 C. diphtheriae causes natural infection only in man. 2 Toxin detection is important to prove the role of C. diphtheriae in causation of disease than just identification of C. diphtheriae. 3 Throat swab should be processed immediately without delay. If any delay is anticipated Amie’s transport medium can be used to transport the specimen. 4 Presumptive identification of C. diphtheriae is made based on the presence of metachromatic granules by Albert’s stain. 5 Loeffler’s serum slope and potassium tellurite agar are the two media frequently used for culture of C. diphtheriae. 6 C. diphtheriae ferments glucose, maltose and sucrose but do not ferment lactose, mannitol and trehalose. 7 C. diphtheriae do not hydrolyse urea and gelatin whereas non-pathogenic C. ulcerans hydrolyses urea.

VIVA 1 Name other staining methods used to demonstrate volutin granules. Ans. Neisser’s and Ponder’s stains. 2 Why serum sugars are used for testing acid production by C. diphtheriae? Ans. These organisms are fastidious. They cannot grow in ordinary sugar media. 3 Name other organisms, which have metachromatic granules. Ans. Bordetella and Brucella.

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FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002

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LESSON

61

Identification of Lactose Fermenting Enterobacteriaceae

LEARNING OBJECTIVES After completing this practical you will be able to 1 Know the procedures used to identify lactose fermenting Enterobacteriaceae and differentiate it from other members of the family Enterobacteriaceae. 2 Perform the tests to identify Escherichia coli and Klebsiella species.

INTRODUCTION Enterobacteriaceae are the part of the normal intestinal flora of animals and humans. They are aerobes and facultative anaerobes. They are non-fastidious and reduce nitrate to nitrites. They are catalase positive and oxidase negative. Based on their action on lactose they are classified into lactose fermenters and non-lactose fermenters. Many of commensal floras of the intestine are lactose fermenters. Late lactose fermenting bacteria are called as para colon bacilli and with the exception of Shigella sonnei all others are commensals. These organisms cause gastrointestinal infection, urinary tract infection, pneumonia and septicaemia. Lactose fermenters such as E. coli, Enterobacter, Klebsiella, Citrobacter and enterotoxigenic E. coli cause watery diarrhoea. The tests used for identification of E. coli and Klebsiella species are listed in the boxes 61-1 and 61-2 respectively.

Gram’s stain Gram stain is indicated in wound infection where pus ells are detected along with Gram negative bacilli. Both centrifuged and un centrifuged urine are subjected for microscopic examination for the detection of pus cells.

2 Culture Stool specimen is inoculated onto MacConkey agar and CLED media and wound swabs are processed using blood agar and MacConkey agar. Inoculated plates are kept for aerobic incubation at 37 °C for 24 hours. Pink colored lactose fermenting colonies of E.coli and K.pneumoniae appear on MacConkey agar plates (Fig. 61-1 and 61-2). On CLED medium E.coli produces lactose fermenting yellow colonies (Fig. 61-3). These are subjected to testing by various biochemical tests to identify the bacteria.

3 Biochemical tests Biochemical tests and other tests, as summarized in the tables are performed to identify E. coli (Fig. 61-4) and Klebsiella species (Fig. 61-5).

4 Antibiotic susceptibility testing Antibiotic susceptibility testing is carried out for the identified bacteria which provide the important information of the susceptibility pattern of the organism.

SPECIMENS Stool, urine, wound swab, sputum, etc. In this chapter, following tests will be described which are routinely used for identification of E.coli and Klebsiella spp.

TESTS FOR IDENTIFICATION OF E. COLI AND KLEBSIELLA SPP. 1 Direct examination

FIGURE 61-1 Mc Conkey agar with lactose fermenting pink colonies of Escherichia coli.

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–

–

FIGURE 61-4 IMViC test for Escherichia coli.

– FIGURE 61-3 CLED Medium with lactose fermenting yellow colonies of Escherichia coli.

+

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–

+

+

FIGURE 61-5 IMViC test for Klebsiella pneumoniae.

BOX 61-1 IDENTIFICATION OF ESCHERICHIA COLI

BOX 61-2 IDENTIFICATION OF KLEBSIELLA SPECIES

Gram stain Gram negative bacilli.

Gram stain Short thick gram negative bacilli.

Motility by hanging drop preparation method Motile.

Motility by hanging drop preparation method Non motile.

Biochemical tests Catalase test: Positive. Oxidase test: Negative. Nitrate reduction test: Reduced to Nitrite. Kligler’s iron agar medium: Acid/Acid. No H2S gas is produced. Fermentation of sugars (glucose, lactose and mannitol): Acid and gas. Methyl red test: Positive. Voges-Proskauer test: Negative. Indole test: Positive. Citrate utilization test: Negative. Urease test: Negative. Lysine decarboxylation test: Positive.

Biochemical tests Catalase test: Positive. Oxidase test: Negative. Nitrate reduction test: Reduced to Nitrite. Kligler’s iron agar medium: Acid/Acid. gas is present. Fermentation of sugars (glucose, lactose and mannitol): Acid and gas. Methyl red test: Negative. Voges-Proskauer test: Positive. Indole test: Negative. Citrate utilization test: Positive. Urease test: Positive (only K. pneumoniae). Lysine decarboxylation test: Positive.

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KEY FACTS 1 Enterobacteriaceae is the part of the normal intestinal flora of animals and humans. 2 Lactose fermenters such as Enterobacter, Klebsiella and Citrobacter and enterotoxigenic E. coli causes watery diarrhoea. 3 Antibiotic susceptibility testing should be done for the identified organism which will give the important information of the susceptibility pattern of the organism.

VIVA 1 Name the indicator used in MacConkey agar medium? Ans. Neutral red. 2 Why some bacteria are late lactose fermenters? Ans. They lack the enzyme lactose permease. 3 What is the color of the colonies of lactose fermenting organism on CLED medium? Ans. Yellow colour. 4 Lactose fermenting colonies grown on MacConkey agar plate are not picked up for oxidase test why? Ans. Acidic pH of the colonies will give false positive reactions. 5 Differentiate between E. coli and Klebsiella species.

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002

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LESSON

62

Identification of Vibrio cholerae

LEARNING OBJECTIVES After completing this practical you will be able to 1 Know about the clinical importance of Vibrio cholerae. 2 Perform the tests to identify V. cholerae and differentiate it from other Vibrio species.

INTRODUCTION V. cholerae are short curved or straight Gram negative bacilli measuring about 1.5 µ × 0.2 to 0.4 µ. These organisms are actively motile by means of a single sheathed polar flagellum. They are strongly aerobic and they grow in alkaline pH (7.4 to 9.6) media. They cause cholera, a toxin mediated disease. Cholera occurs in many forms sporadic, endemic, epidemic or pandemic. Cholera is an exclusively human disease. Infections are spread by feco-oral transmission. List of tests for identification of V.cholerae is summarized in the box 62-1.

the corners of cover slip. A loopful of stool specimen is collected by a heat sterilised wire loop and transferred it on to the middle of the cover slip. This is placed over a clean cavity slide in such a way that the drop hangs from the cover slip to cavity of the slide. This preparation is observed first under 10x followed by 40x, for the presence of characteristic darting motility of V.chloerae. Observation of darting motility in stool sample helps in presumptive diagnosis of cholera, which should be confirmed by biochemical and serologic methods.

2 Culture Alkaline peptone water is used as enrichment medium. DCA and TCBS are employed as the selective media for V. cholerae. A loopful of stool is inoculated into alkaline peptone water and incubated at 37°C for 6 hours. Then this is sub cultured on to TCBS medium, and incubated at TCBS at 37°C overnight. After overnight incubation the presence of yellow colonies on TCBS medium and non lactose fermenting colorless colonies on DCA is looked for. The biochemical tests of these colonies are performed for identification of V. cholerae.

SPECIMENS 3 Biochemical tests Stool (rice watery stool) and rectal swab. In this chapter the following tests will be described which are employed routinely for identification of V. cholerae.

TESTS FOR IDENTIFICATION OF VIBRIO CHOLERAE 1 Direct examination Hanging drop preparation This test is carried out to demonstrate motility of V.chloerae, which are actively motile. A clean cover slip is taken and vaseline is applied over all

Biochemical tests such as lysine and ornithine decarboxylase and arginine dihydrolase (Fig. 62-1) and other tests performed to identify V. cholerae are summarized in the box 62-1. Biochemical tests like Voges Proskauer, polymyxin-B (50 U) sensitivity (Fig. 62-2), sheep RBC hemolysis and chick cell agglutination are done further for the differentiation of two biotypes Classical and Eltor (Table 62-1).

4 Antibiotic susceptibility testing Antibiotic susceptibility testing is carried out for the identified bacteria which provide the important information of the susceptibility pattern of the organism.

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BOX 62-1 IDENTIFICATION OF VIBRIO CHOLERAE Gram stain Gram negative curved or straight rods. Motility by hanging drop preparation method Actively motile. Biochemical tests Catalase test: Positive.

+

+

–

FIGURE 62-1 Lysine and ornithine decarboxylase and arginine dihydrolase tests for Vibrio cholerae.

Oxidase test: Positive. Nitrate reduction test: Reduced to Nitrite. Kligler’s iron agar medium – K/A. Fermentation of sugars (glucose, sucrose and mannose fermented: Acid only, lactose not fermented). Indole test: Positive. Citrate utilization test: Positive. Urease test: Negative. Lysine decarboxylation test: Positive. Ornithine decarboxylation test: Positive. Arginine dihydrolase test: Negative. Confirmation of V. cholerae isolates is done by serotyping with specific O antisera (slide agglutination test) using O1 Ogawa and, O1 Inaba antisera.

FIGURE 62-2 Polymyxin B sensitivity of Vibrio cholerae.

Table62-1 Differences between V. cholerae biotype classical and V. cholerae biotype El Tor Test Voges-Proskauer (VP) test Sheep RBCs haemolysis Chick RBCs agglutination Polymyxin B sensitivity Susceptibility to Mukherjee’s phage IV Sensitivity to Vibriostatic (O/129) agent Susceptibility to group V phage

V. cholerae biotype classical

V. cholerae biotype El Tor

+

+ + + -

+

-

+ -

+

KEY FACTS 1 Subculture from the incubated alkaline peptone water to be done within 6 hours of incubation. 2 Looking for the presence of darting motility is not recommended for the diagnosis of cholera because this type of motility can also be observed in other bacteria. 3 Biochemical tests like Voges Proskauer, polymyxin B sensitivity, sheep RBC hemolysis, chick cell agglutination are usually carried out to differentiate between V. cholerae biotypes Classical and Eltor.

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VIVA 1 What are non agglutinable vibrios? 2 Mention the application of vibriostatic O/129 reagent. 3 Why yellow colonies are produced on TCBS while culturing V. cholerae? Ans. Sucrose fermentation degreases pH, which converts bromo thymol blue to yellow colour. 4 What is the chemical agent used for string test for V. cholerae? Ans. Sodium deoxycholate (0.5 %). 5 How the haemodigestion of V. cholerae differs from hemolysis of other bacteria? Ans. Haemodigestion is not toxin mediated. It is mediated by peroxides.

FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002.

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63

Identification of Pseudomonas aeruginosa

LEARNING OBJECTIVES

2 Culture

After completing this practical you will be able to 1 Know the clinical importance of Pseudomonas aeruginosa. 2 Perform the tests to identify P. aeruginosa.

Pus swab is inoculated on to blood agar, MacConkey agar and nutrient agar and the plates are incubated at 37°C overnight. Moist hemolytic colonies are produced on blood agar (Fig. 63-1). Greenish blue pigmented colonies are produced on nutrient agar (Fig. 63-2). P. aeruginosa colonies produce earthy or grape juice like odour.

INTRODUCTION P. aeruginosa are non-fermentative slender Gram negative bacilli that measure about 1.5 µm to 3µm by 0.5 µm in size. They are obligate aerobes. They grow at a temperature range of 6°C to 42°C. They produce opaque irregular colonies with earthy smell and form non-lactose fermenting colonies on MacConkey agar. In liquid media they form a surface pellicle. P. aeruginosa produces pigments such as pyocyanin, pyoverdin and pyorubrin. They utilise carbohydrates oxidatively. Pseudomonas causes infection of burns and wounds and is a common cause of nosocomial infections. The tests for identification of P. aeruginosa are mentioned in the box 63-1.

SPECIMENS Pus swab. In this chapter the following tests will be described which are employed routinely for identification of P. aeruginosa.

TESTS FOR IDENTIFICATION OF PSEUDOMONAS AERUGINOSA 1 Direct examination Gram stain Gram stain of the pus swab reveal thick Gram negative bacilli measuring 1.5–3 µm by 0.5 µm in size.

3 Oxidase test P. aeruginosa is an oxidase positive bacterium. It produces the enzyme cytochrome oxidase. The enzyme in the presence of oxygen oxidizes 1% tetramethyl paraphenylene diamine dihydrochloride and produces a deep purple colour. A filter paper strip impregnated with 1% tetramethyl paraphenylene diamine dihydrochloride is taken. By a platinum wire, the colonies on nutrient agar are picked up and streaked on the surface of the filter paper. Positive test is indicated by appearance of deep purple color, and negative test by no color change (refer chapter 21). Pseudomonas and other nonfermentative Gram negative bacilli are oxidase positive whereas all Enterobacteriaceae bacteria are oxidase negative.

4 Biochemical tests They are non-fermenters. They break down glucose oxidatively with production of acid only (Fig. 63-3).

5 Antibiotic susceptibility testing Antibiotic susceptibility testing is carried out for the identified bacteria which provide the important information of the susceptibility pattern of the organism (refer chapter 31).

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BOX 63-1 IDENTIFICATION OF PSEUDOMONAS AERUGINOSA Gram stain Gram negative bacilli. Motility Motile. Biochemical tests Growth at 42°C: Positive. Catalase test: Positive. Oxidase test: Positive. Nitrate reduction test: Reduced to nitrite. Kligler’s iron agar median: K/K Oxidative breakdown of sugars glucose: acid only, lactose, mannitol, sucrose, maltose: No acid. Indole test: Negative. Citrate utilization test: Positive. Urease test: Variable. Lysine decarboxylation test: Negative. Arginine dihydrolase test: Positive.

FIGURE 63-2 Pseudomonas colonies on nutrient agar.

– FIGURE 63-1 Non hemolytic Pseudomonas colonies on blood agar.

+

FIGURE 63-3 OF test showing oxidative utilization of glucose.

KEY FACTS 1 2 3 4

P. aeruginosa are non-fermentative slender Gram negative bacilli. They utilise carbohydrates oxidatively. P. aeruginosa produces greenish blue pigmented colonies on nutrient agar. P. aeruginosa is an oxidase positive bacteria.

VIVA 1 Name the selective media used for the isolation of P. aeruginosa. Ans. Cetrimide agar. 2 Name different pigments produced by P. aeruginosa? FURTHER READINGS 1 2 3 4 5

Collins CH, Lyne PM and Grange JM. Microbiological Methods. Butterworths, London, 94-96, 1995. Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Jawetz, Melnick and Adelberg. Medical Microbiology. 23rd Edition. McGaw Hill. 2003. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. 1996. Murray PR, Rosenthal KS, Kobayashi GS and Pfaller ME. Medical Microbiology. 5th Edition. Mosby Inc. 2002.

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UNIT

IX Parasitology

Introduction Lesson 64 Saline Wet Mount of Stool Lesson 65

Iodine Wet Mount of Stool

Lesson 66

LPCB Wet Mount of Stool

Lesson 67 Acid-fast Staining of Stool Smears Lesson 68 Leishman’s Staining of Peripheral Blood Smears Lesson 69 Concentration of Stool for Parasites Lesson 70 Culture of Stool for Entamoeba histolytica

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Introduction Parasitology traditionally includes the study of three major groups of animal parasites: parasitic protozoa, parasitic helminths (worms), and those arthropods that directly cause disease or act as vectors of various pathogens. Infections of humans caused by parasites number in the billions and range from relatively innocuous to fatal. The diseases caused by parasites constitute major human health problems throughout the world. The incidence of many parasitic diseases (e.g. schistosomiasis, malaria) have increased rather than decreased in recent years. Other parasitic illnesses have increased in importance as a result of the AIDS epidemic (e.g., cryptosporidiosis, Pneumocystis carinii pneumonia, and strongyloidiasis). The unicellular parasites (protozoa) and multicellular parasites (helminths, arthropods) are antigenically and biochemically complex, as are their life histories and the pathogenesis of the diseases they cause. The basis for effective treatment and cure of a patient is the rapid diagnosis of the disease and its causative agent, which is based on the analysis of the clinical symptoms in combination with laboratory tests. As we are in the 21st century, clinicians are becoming increasingly able to diagnose and treat diseases at the molecular level. During past few years, there has been an increased awareness of the importance of trained and qualified personnel to perform diagnostic procedures. It becomes even more important to provide well-written lab protocols and to standardize test methods for consistency. Therapy based on patient history and symptoms is not generally recommended in cases of parasitic infections. Thus understanding of characteristics of any parasitic infection and the use of appropriate diagnostic procedures accompanied by a complete understanding of the limitations of each procedure become very important.

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LESSON

64

Saline Wet Mount of Stool

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform saline wet mount preparation of faeces for demonstration of intestinal parasites. 2 Identify intestinal protozoal cysts, trophozoites, helminthic eggs and larva based on the recognition of specific morphological characters, in the stool specimen.

II Reagents and glass wares Microscopic slides, cover slips, physiological saline, and applicator stick Preparation of physiological saline: Normal saline is also called physiological saline. It is 0.85% sodium chloride in distilled water. It is prepared by weighing 8.5 gm of sodium chloride and dissolving it in 1000 ml of distilled water. III Specimen Fresh stool specimen is required for stool microscopy.

INTRODUCTION Stool microscopy is an easy and rapid method employed for detection of intestinal ova and cysts with the aid of wet mount preparations. The wet mount preparations most commonly used for detection are the i) saline wet mount preparation and ii) iodine wet mount preparation. Of late, lacto-phenol cotton blue (LPCB) wet mount preparation is also used. The wet mount preparations of stool specimen are screened first under the low power objective (10x) and then under high power (40x). The saline preparation is mainly used to a. demonstrate the presence of trophozoites by demonstrating the motility and b differentiate between bile stained and non bile-stained eggs.

PROCEDURE 1 Take a clean glass slide. 2 With help of a glass dropper, put a drop of saline on the glass slide. 3 With the help of an applicator stick, take a small portion of the stool specimen (match stick head size) and emulsify it in the drop of saline. 4 Put a cover slip over the saline suspension of the stool. 5 Examine the preparation first under the low power (10x) and then under high power (40x) objective of the microscope. 6 Record the findings with description of the morphological characteristics.

PRINCIPLE QUALITY CONTROL

The saline wet mount preparation is prepared by mixing a small quantity of faeces with physiological saline. The saline wet mount is a colourless preparation that highlights the staining property of the egg, whether bile or non-bile stained, detects motility of trophozoites, and facilitates demonstration of chromatoidal bodies in the cyst.

A saline and iodine wet mount preparation of known positive stool specimen for protozoal cysts, larva and helminthic eggs are prepared. These preparations are observed and compared with the saline and iodine wet mount preparations of the test stool specimen for various morphological forms of the parasite and identified.

REQUIREMENT

OBSERVATIONS

I Equipments Compound light microscope.

1 A 10–15 µm sized round refractile, colourless structure seen with four nuclei, though not distinct.

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2 A 15–30µm sized, round refractile, colourless structure seen with at least eight nuclei, though not distinct. 3 A 6–10µm sized oval shaped colourless structure seen with distinct wall and faint axostyle. 4 A 60µm × 40µm sized, oval shaped, non-bile stained, colourless structure seen with transparent hyaline shell membrane and segmented ovum with blastomeres. A clear space is visible between the thin hyaline shell membrane and blastomeres. 5 A 60–75µm × 40–50µm sized, round oval shaped, bile stained structure seen with thick translucent shell with an albuminous coat. It contains a very large conspicuous unsegmented ovum with a clear space at each pole. 6 A 25µm × 50µm sized, bile coloured, and barrel shaped structure seen with visible mucus plugs at each pole. It has a double layered egg shell enclosing a visible unsegmented ovum.

3 4 5 6

Cyst of Giardia intestinalis (Fig. 64-2). Hook worm egg (Fig. 64-3). Fertilized egg of Ascaris lumbricoides (Fig. 64-4). Egg of Trichuris trichiura. (Fig. 64-5).

RESULTS AND INTERPRETATION 1 Cyst of Entamoeba histolytica/dispar (Fig. 64-1). 2 Cyst of Entamoeba coli.

FIGURE 64 -3 Hook worm egg, x 400.

FIGURE 64 -1 Cyst of Entamoeba histolytica/dispar, x 400.

FIGURE 64 -4 Fertilized egg of Ascaris lumbricoides, x 100.

FIGURE 64 -2 Cyst of Giardia intestinalis, x 400.

FIGURE 64 -5 Eggs of Trichuris trichiura, x 400.

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KEY FACTS 1 A stool wet mount preparation should always first be screened under the low power objective (10x) and then under the high power (40x) for identification. 2 A wet mount preparation should neither be too thick nor thin. The preparation should be such that, a printed letter should be read through it. The wet mount preparation must not overflow. 3 Trophozoites and larvae are visualised best in the saline wet mount. 4 Bile staining property of the parasitic egg can be appreciated.

VIVA 1 List the normal constituents of stool. Ans. The normal constituents of stool are yeast cells, cotton fiber, starch cell, potato parenchymal cell, plant epidermal hairs, fungal spores, muscle fibres, vascular structure of plants, pollen, stone cell, plant cells, oil drops, rice starch, potato starch, corn starch, leucocytes, moulds and bacteria. 2 What are the different wet mount preparations of stool? Ans. Different wet mount preparations of stool are saline wet mount, iodine wet mount, LPCB wet mount, and buffered methylene blue. 3 What are the uses of saline wet mount?

FURTHER READINGS 1 Garcia LS. Diagnostic Medical Parasitology. ASM press, Washington D.C. 4th Edition. 2003. 2 Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3rd Edition. 2006. 3 Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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LESSON

65

Iodine Wet Mount of Stool

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform saline wet mount preparation of faeces for demonstration of intestinal parasites. 2 Identify intestinal protozoan cysts, trophozoites, helminthic eggs and larva based on the recognition of specific morphological characters, in the stool specimen by iodine wet mount.

INTRODUCTION Iodine wet mount is prepared by using a drop of saline for preparation of wet mount of stool. Different types of iodine solution are used. These are Dobell and O’Connor’s iodine, Lugol’s’ iodine and D’ Antonie’s iodine. Iodine wet mount is mainly used for protozoal cysts. The preparation clearly demonstrate the presence of nuclei in protozoan cyst as brown dots, and also demonstrates the presence of glycogen mass in protozoan cyst.

PRINCIPLE The iodine wet mount preparation is a brown coloured preparation that highlights the presence of pale refractile nuclei, yellowish cytoplasm and brown glycogen material within the cysts. Bile staining property of the helminthic eggs cannot be appreciated in the iodine preparation, as it is already coloured. The motility of trophozoites is also inhibited in the iodine wet mount. List of bile-stained and non-bile stained eggs are provided in the table 65-1.

REQUIREMENTS I Equipments Compound light microscope.

II Reagents and glass wares Microscopic slides, cover slips, applicator stick and Dobell and O’Connor’s iodine. Preparation of Dobell’s iodine: Dobell’s iodine is used for making iodine wet mount of stool. It contains iodine and potassium iodide in distilled water. It is prepared by weighing 2 grams of iodine crystals and dissolving it in 100 ml of distilled water. Then 4 grams of potassium iodide is weighed and added to the solution prepared. It is mixed well and Dobell’s iodine solution is prepared. III Specimen Fresh stool specimen is required for stool microscopy.

PROCEDURE 1 Take a clean glass slide. 2 With the help of a glass dropper, put a drop of iodine on the glass slide. 3 With the help of an applicator stick, take a small portion of the stool specimen (match stick head size) and emulsify it in the drop of iodine. 4 Put a cover slip over the iodine suspension of the stool. 5 Examine the preparation first under the low power (10x) and then under high power (40x) objective of the microscope. 6 Record the findings with description of the morphological characteristics.

QUALITY CONTROL A saline and iodine wet mount preparation of known positive stool specimen for protozoal cysts, larva and helminthic eggs are prepared. These preparations are observed and compared with the saline and iodine wet mount preparation of the test stool specimen for various morphological forms of the parasite and identified.

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OBSERVATIONS 1 A 10–15µm sized, round, yellow coloured, structure seen with brown coloured 1–4 nuclei and glycogen mass. 2 A 15–30µm sized, round, yellow coloured structure seen with brown coloured 1–8 nuclei and diffuse glycogen mass. 3 A 6–10µm sized oval shaped brown coloured structure seen with visible axostyle and distinct cyst wall surrounded by a halo. 4 A 60µm × 40µm sized, oval shaped, yellow coloured structure seen with lightly stained shell membrane and segmented ovum with light yellow stained blastomeres. A clear space is visible between the stained shell membrane and blastomeres. 5 A 60–75µm×40–50µm sized, round oval shaped, yellow coloured structure seen with yellow stained outer corticated thick cell wall. The unsegmented ovum and also the space between the shell and ovum at each pole are stained yellow. 6 A 25µm × 50µm sized yellow coloured, barrel-shaped structure seen with lightly stained mucus plugs at each pole. The egg shell is stained brown and encloses the light yellow stained unsegmented ovum.

FIGURE 65-2 Cyst of Giardia intestinalis, x 400.

RESULTS AND INTERPRETATION 1 2 3 4 5 6

Cyst of Entamoeba histolytica/dispar (Fig. 65-1). Cyst of Entamoeba coli. Cyst of Giardia intestinalis (Fig. 65-2). Hook worm egg (Fig. 65-3). Fertilized egg of Ascaris lumbricoides (Fig. 65-4). Eggs of Trichuris trichiura (Fig. 65-5).

FIGURE 65-3 Hook worm egg, x 400.

Table 65-1 List of bile-stained and non-bile stained eggs Bile stained eggs Ascaris lumbricoides, Trichuris trichiura, Taenia species, Echinococcus species, Diphyllobothrium latum, Schistosoma mansoni, Schistosoma japonicum, Schistosoma hematobium, Fasciola hepatica, Fasciolopsis buski, Paragonimus westermani, and Clonorchis sinensis.

FIGURE 65- 1 Cyst of Entamoeba histolytica/dispar, x 400.

Non-bile stained eggs Hook worm, Enterobius vermicularis and Hymenolepis nana.

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FIGURE 65- 4 Fertilized egg of Ascaris lumbricoides, x 400.

FIGURE 65 –5 Eggs of Trichuris trichiura, x 400.

KEY FACTS 1 2 3 4 5

A stool wet mount should always be screened immediately. A wet mount preparation should neither be too thick or thin. Iodine wet mount kills both living trophozoites of protozoa and larvae of worms. Bile staining property of the parasitic egg cannot be appreciated in the iodine preparation as it is already coloured. Iodine solution should always be handled with care, as it proves injurious if inhaled or comes in contact with eyes.

VIVA 1 What are the uses of iodine wet mount? 2 What are the different types of iodine solution that can be used for stool wet mount preparation? 3 List the advantages and disadvantages of iodine wet mount. Ans. Advantages a The number of nuclei present in the protozoan cyst can be clearly distinguished. b It also demonstrates the yellowish cytoplasm and brown glycogen mass present in the cysts. Disadvantages a Trophozoites are killed by iodine, hence cannot be demonstrated in iodine wet maint. b The bile staining property of the helminthic eggs cannot be made out. c Chromatoidal bars in protozoan cysts are not clearly demonstrable. 4 List bile stained eggs. 5 List non-bile stained eggs.

FURTHER READINGS 1 Garcia LS. Diagnostic Medical Parasitology. ASM press, Washington D.C. 4th Edition. 2003. 2 Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3rd Edition. 2006. 3 Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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LESSON

66

LPCB Wet Mount of Stool

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to:

I Equipments Compound light microscope.

1 Perform lacto-phenol cotton blue (LPCB) wet mount preparation of faeces for demonstration of intestinal parasites. 2 Identify intestinal protozoal cysts, trophozoites, helminthic eggs and larva based on the recognition of specific morphological characters, in the stool specimen by the LPCB wet mount

II Reagents and glass wares Microscopic slides, cover slips, applicator stick and lactophenol cotton blue (LPCB) reagent. Preparation of LPCB solution: Weigh 20 gm phenol crystals, and transfer it to a beaker (100 ml capacity). Measure 20 ml of distilled water in a measuring cylinder and add to the phenol, and mix it well. Measure 20 ml of lactic acid. Then transfer to the bottle. Measure 40 ml of glycerol and then transfer to the bottle. Mix it well. Dissolve the solution by heating gently over a spirit flame. Then allow it to be cooled. Weigh 0.05 gm of cotton blue. Add this to the solution and mix it well. Transfer the LPCB solution to a clean, leak proof, brown bottle. Label the bottle and mark it. For daily use, students can transfer about 10 ml of LPCB solution to a small brown dropper bottle or insert a dropping pipette through the cap of a small brown bottle.

INTRODUCTION Lacto-phenol cotton blue (LPCB) is a staining reagent which is extensively used in the examination of clinical specimens for demonstration of fungi and fungal elements. Of late, the LPCB has also been used in the direct wet mount preparation of stool specimens for demonstration of larvae, ova and cysts of the intestinal parasites.

III Specimen Fresh stool specimen is required for stool microscopy.

PRINCIPLE The LPCB is a combined fixative, staining and clearing agent. It contains cotton blue which stains both the helminthic ova and protozoal cysts deep blue. It contains phenol and lactic acid which clears faecal debris. Glycerol in the LPCB provides a semi-permanent preparation. Therefore, in the wet mount preparation of LPCB, blue-colour stained cysts of intestinal protozoa and ova of eggs could easily be detected and identified. Helminthic ova are stained such a deep blue that it is difficult to miss them even during screening with a low power objective. Additional advantage of the LPCB is that it can also detect blue coloured Cyclospora and Isospora, the intestinal coccidian parasites, in the stool. Vegetable cells, mucus, muscle fibres and other artifacts are clearly stained with LPCB stain. They are still recognisable as artifacts when the LPCB stain is used.

PROCEDURE 1 Take a clean glass slide. 2 With help of a glass dropper, put a drop of LPCB on the glass slide. 3 With the help of an applicator stick, take a small portion of the stool specimen (match stick head size) and emulsify it in the drop of LPCB. 4 Put a cover slip over the LPCB suspension of the stool. 5 Examine the preparation first under the low power (10x) and then under high power (40x) objective of the microscope. Note: Examine at least 30 minutes after preparation of the wet mount. 6 Record the findings with description of the morphological characteristics.

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QUALITY CONTROL

5 Fertilized egg of Ascaris lumbricoides (Fig. 66-4). 6 Egg of Trichuris trichiura.

A LPCB wet mount preparation of known positive stool specimen for protozoal cysts, larva and helminthic eggs are prepared. These preparations are observed and compared with the LPCB wet mount preparation of the test stool specimen for various morphological forms of the parasite and identified.

OBSERVATION 1 A 10–15 µm sized, round, deep blue coloured, structure seen with blue coloured 1–4 nuclei and glycogen mass. 2 A 15–30 µm sized, round, deep blue coloured structure seen with blue coloured 1–8 nuclei and diffuse glycogen mass. 3 A 6–10 µm sized oval shaped deep blue coloured structure seen with visible axostyle and distinct cyst wall surrounded by a halo. 4 A 60µm × 40µm sized, oval shaped, deep blue coloured structure seen with lightly stained shell membrane and segmented ovum with deep blue stained blastomeres. A clear space is visible between the stained shell membrane and blastomeres. 5 A 60–75 µm × 40–50µm sized, round oval shaped, deep blue coloured structure seen with blue stained outer corticated thick cell wall. The unsegmented ovum and also the space between the shell and ovum at each pole are stained blue. 6 A 25µm × 50µm sized blue coloured, barrel -shaped structure seen with lightly stained mucus plugs at each pole. The egg shell is stained blue and encloses the blue stained unsegmented ovum.

FIGURE 66 - 2 Cyst of Giardia intestinalis, x 400.

RESULTS AND INTERPRETATION 1 2 3 4

Cyst of Entamoeba histolytica/dispar (Fig. 66-1). Cyst of Entamoeba coli. Cyst of Giardia intestinalis (Fig. 66-2). Hook worm egg (Fig. 66-3).

FIGURE 66 - 1 Cyst of E. histolytica / dispar, x 400.

FIGURE 66 - 3 Hook worm egg, x 400.

FIGURE 66 - 4 Fertilzed egg of Ascaris lumbricoides, x 100.

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KEY FACTS 1 2 3 4

LPCB wet mount of stool is always examined at least 30 minutes after preparation of the wet mount. A wet mount preparation should neither be too thick nor thin. LPCB kills the trophozoites of Entamoeba, Giardia and Trichomonas, hence can not be demonstrated by LPCB. In LPCB preparation, both bile-stained and non bile-stained helminthic eggs are stained blue.

VIVA 1 What are the uses of LPCB mount? 2 List the advantages and disadvantages of the iodine wet mount.

FURTHER READINGS 1 Garcia LS. Diagnostic Medical Parasitology. ASM press, Washington D.C. 4th Edition. 2003. 2 Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3rd Edition. 2006. 3 Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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LESSON

67

Acid-fast Staining of Stool Smears

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform modified acid fast staining of faeces for demonstration of intestinal coccidian parasites. 2 Detect and recognize the presence of acid fast oocysts in the stool smear stained by acid-fast method.

II Reagents and lab wares Inoculation loop, carbol fuchsin, 5% aqueous sulphuric acid (decolouriser), and 0.3% methylene blue (counter stain). III Specimen Stool specimen.

PROCEDURE INTRODUCTION Intestinal coccidian parasites, namely Cryptosporidium parvum, Cyclospora cayetanensis and Isospora belli cause infection of the gastrointestinal tract of humans. The infection causes diarrhoea, which may be serious in the immunocompromised patients. Oocysts, the diagnostic morphological form of the parasite, are usually excreted in the human faeces. These oocysts are demonstrated by the modified acid fast staining technique as pink-coloured acid fast structures against a blue background.

PRINCIPLE The principle of modified acid fast staining is based on the fact that the oocyst of these coccidian parasites are acid fast and retain the basic dye (dilute carbol fuchsin)appearing pink. They do not get decolourised with the acid alcohol and hence do not take up the counter stain methylene blue. The stool material in the background gets decolourised easily and take up the counter stain appearing blue. Both hot and cool modified acid stains have been used with equal sensitivity. Acid fast parasites and parasitic components are listed in the box 67-1. In this chapter hot modified acid fast technique will be described.

REQUIREMENTS I Equipments Compound light microscope and Bunsen flame.

1 Make a smear of stool on a clean glass slide. 2 Heat fix the smears by heating at 70°C for 10 minutes. 3 Put the smears on a slide rack and flood the smear with carbol fuchsin. 4 Heat the slides from below intermittently by Bunsen flame until the steam rises. Do not allow the stain to dry on the slide, and if necessary add more carbol fuchsin to cover the smear. The slide is allowed to stain for 9 minutes. 5 Wash the smears with tap or distilled water. 6 Cover the smear with 5% aqueous sulphuric acid, as decolouriser, for 30 seconds. 7 Wash the slides with water to remove all traces of acid. 8 Cover the smear with methylene blue, as counter stain, for 1 minute. 9 Rinse the smears again under tap water and air dry it. 10 Observe the smear first under low power (10x) objective, and then under oil immersion (100x) objective. Note: The smear should be examined following a zig-zag pattern for at least 10–30 minutes, before declaring the smear negatives. 11 Record the observations in the note book. Findings are recorded, together with grading of the positive smear.

QUALITY CONTROL A known positive control stool smear stained by the modified acid fast staining method containing the pink coloured acid fast oocysts of C. parvum. The stained smear is compared with the control smear for appropriate oocyst morphology and staining appearance.

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BOX 67-1 ACID FAST PARASITES AND PARASITIC COMPONENTS The intestinal coccidian parasites Cryptosporidium parvum. Cyclospora cayetanensis. Isospora belli. The non- intestinal coccidian parasites Toxoplasma gondii. Sarcocystis hominis. Other parasitic components Scolices of Echinococcus granulosus. Spores of Microsporidia. Eggs of Taenia saginata.

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RESULTS AND INTERPRETATION 1 The acid fast structure is oocyst of C. parvum (Fig. 67-1). 2 The acid fast structure is the oocyst of I. belli (Fig. 67-2). 3 The acid fast structure is the oocyst of C. cayetanensis (Fig. 67-3). Note: Cyclospora oocysts are variably acid fast.

OBSERVATIONS 1 Red coloured spherical acid fast structure measuring 4-6µm diameter seen against a blue background of stool smear. 2 Red coloured elliptical shaped acid fast structure measuring 20µ m x 10µm seen against a blue background of stool smear. 3 Red coloured, round to ovoid shaped acid fast structure measuring 8µm–10µm diameter seen against a blue background of stool smear. Some structures are acid fast while others are not.

FIGURE 67- 1 Oocyst of C. parvum, x 1000.

FIGURE 67- 2 Oocyst of I. belli, x 1000.

FIGURE 67- 3 Oocyst of C. cayatenensis, x 1000.

KEY FACTS 1 Modified acid fast staining is a widely used method for the demonstration of the oocysts of intestinal coccidian parasites in stool. 2 It is an example of permanent staining of stool. 3 Stool specimens should be handled carefully, as infection is caused by ingestion of oocysts from hands. 4 Oocysts of Cryptosporidium and Isospora are consistently acid fast in nature while Cyclospora is variably acid fast. 5 Cyclospora oocysts are twice the size of the Cryptosporidium oocysts. 6 Cryptosporidium oocysts are to be distinguished from yeast cells which are non acid fast and show budding and variable sizes.

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VIVA 1 List the intestinal and non intestinal coccidian parasites. 2 How are oocysts of Cryptosporidium differentiated from yeast cells in this staining technique? Ans. The features that differentiate the oocyst of Cryptosporidium from that of yeast cells in modified acid-fast staining are that oocyst of Cryptosporidium are acid-fast while that of the yeast cells are non-acid fast and are of variable sizes with budding. 3 Name the coccidian parasite associated with traveler’s diarrhoea. Ans. Cyclospora cayetanensis and Cryptosporidium parvum. 4 Name the stool concentration method for intestinal coccidian parasites? Ans. Sheather’s sucrose floatation method. 5 List all acid fast parasitic components.

FURTHER READINGS 1 Garcia LS. Diagnostic Medical Parasitology. ASM press, Washington D.C. 4th Edition. 2003. 2 Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3rd Edition. 2006. 3 Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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LESSON

68

Leishman’s Staining of Peripheral Blood Smears

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform Leishman’s staining of peripheral blood smear for malaria parasites and microfilariae. 2 Detect and identify important parasites of blood such as malarial parasites (various morphological forms) and microfilaria of filarial worm.

INTRODUCTION Haemoparasites are found in the peripheral blood smears of many parasitic diseases. These include i). protozoal infections such as malaria, babesiosis, Chagas’ disease, African trypanosomiasis and leishmaniasis; and ii) helminthic infections such as lymphatic filariasis, and loiasis. Examination of permanent stained blood smears is essential for detection and specific identification of blood parasites. Thick and thin blood smears stained with Romanowsky’s stains are the permanent staining methods widely used for demonstration of blood parasites. These blood smears, although simple and easy to prepare require experienced eyes for the screening, detection and identification of these haemoparasites. Microfilariae although can be identified in wet mount preparation of fresh blood, on basis of their shape and motility, their correct and accurate identification is possible only after permanent staining of blood smear. For malarial parasites, blood films help quantitate parasitaemia, in addition to species identification.

PRINCIPLE As a rule thin blood smears are fixed by alcohol or other fixatives before staining. Two types of stains are used. One type of stains (e.g. Leishman’s, Wright’s, etc.) has both fixatives and staining reagents. The other type of reagents (e.g. Giemsa, Field, JSB stains) has only staining reagents. Therefore, the thin films need prior fixation before staining with any of these stains.

Thin blood films are used primarily for identification of species of malarial parasites. Thin blood films are of monolayer thickness, allows better identification of species. The disadvantage of thin blood smear is that as less amount of the blood is examined, the number of parasites per field too are much less than that of thick blood smear. The procedure followed for staining thick smears are essentially the same as for thin smears except that the initial steps of fixation smears by methanol is not done. The absence of fixation of the smears by methanol allows lysis of red blood cells and dehaemoglobinisation by aqueous stain solution. Thick blood smear is primarily used as a screening procedure. It is a sensitive procedure for detection of parasites compared to thin smear, as it allows examination of larger amount of blood. Disadvantage of thick blood smear is that species identification of parasites cannot be made. Thick blood smears are several layers thick, allows only detection of parasites. The parasites found in the peripheral blood smear are listed in the box 68-1.

REQUIREMENTS I Equipments Compound light microscope. II Reagents and lab wares Bunsen flame and clean glass slides, Leishman’s stain, EDTA anticoagulated blood, 70% ethyl alcohol, buffered distilled water and methyl alcohol. Preparation of EDTA anticoagulated blood: It is prepared by dissolving 5 grams of EDTA in 100 ml of distilled water. Then 0.4 ml is aliquoted into tubes and the water is evaporated. Blood can then be added to the anticoagulant or 20 mg of dry EDTA too can be added per tube. Then blood can be added to it. Preparation of Leishman’s stain: Leishman’s stain is prepared by dissolving 0.15 gram of Leishman’s dry powder in 100 ml of absolute methyl alcohol in a bottle. The bottle is shaken until the powder is dissolved and allowed to stand for 48 hours with frequent shaking in between.

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III Specimen Fresh blood obtained by finger prick (or) EDTA anticoagulated blood.

b The thin film only must be fixed in absolute methanol before staining. c Both the thick and thin blood films should be stained simultaneously.

PROCEDURE Leishman’s staining Preparation of thin blood smear 1 Take an alcohol cleaned and grease free slide. Note: If old slides are to be used, they should be first cleaned with detergent and then with 70% ethyl alcohol. 2 Place a small drop of fresh blood on the clean microscopic slide, about 1.5 cm from the end of the slide. 3 Touch the drop of blood with the edge of another slide and allow the blood to spread along the edge. 4 Push the second slide or the spreader across at about a 30° angle, along the surface of the horizontal slide to the far end forming a ‘tongue shaped’ thin film. Note: The thin, feathered end should be at least 2 cm long, and occupy the central area of the slide, with free margins on both sides. 5 Allow the thin blood film to dry, and then stain the smear with Leishman’s stain. 7 First screen the film with the low-power objective of the microscope for detection of microfilaria and examine at least two hundred microscopic fields using 100 x objectives for the presence of malarial parasites.

Preparation of thick blood smear 1 Place two or three small drops of fresh blood (without any anticoagulant) on a grease-free alcohol cleaned slide. 2 Spread and mix the drops of blood with the corner of another slide, over an area of about 2 cm in diameter. The mixing is continued for about 30 seconds to prevent formation of any fibrin strands which may conceal the parasites in a stained smear. Note: If blood containing anticoagulant is used, 2 or 3 drops maybe spread over an area of about 1 cm in diameter. 3 Allow the thick blood film to air-dry at room temperature in a dust free area. 4 After drying, lyse the thick blood film with distilled water for dehaemoglobinisation. 5 After the step of dehaemoglobinisation, stain the film with Leishman’s staining. 6 After staining, air-dry the film in a vertical position. 7 Examine the stained smear under oil-immersion objective (100 x) for detection of malarial parasites and microfilaria.

Preparation of combined thick and thin films In field surveys, it may be helpful to prepare slides with both the thick and thin film on the same slides. Note: With this type of preparation: a Sufficient time should be allowed for the thick portion of the smear to dry before staining.

1 Flood the blood smear with 5–10 drops of the Leishman’s stain and allow to stand for 2 minutes. 2 After 2 minutes, dilute the stain by adding twice as many drops of buffered distilled water. Allow the solution to mix well. 3 Allow the solution to stand for 15–20 minutes. 4 Wash the slide with buffered distilled water, and rinse it dry. 5 Examine the stained slide under the oil immersion objective (100 x) of the microscope.

QUALITY CONTROL A known positive control, Leishman stained thick and thin blood films for malarial parasites and microfilaria, is compared with the stained test blood films for similar morphology of the parasites.

OBSERVATIONS 1 The blood film shows the presence of RBC’s identified as pale round cells with dense periphery and lighter core. Some of the normal sized RBC’s show blue ring-shaped parasite cytoplasm surrounding a central vacuole with a red coloured nucleus (1 chromatin dot) present at its centre. Often 2 or more ring forms of the parasite are found inside a single RBC. a Neutrophils in the blood films are identified by their multilobed nucleus. b Lymphocytes are identified as having big nucleus pushing the cytoplasm to the periphery. c Monocytes have kidney shaped nucleus. 2 The blood film shows the presence of normal RBC’s, neutrophils, lymphocytes and monocytes. In addition, typical crescent (banana) shaped structures are seen with dark blue cytoplasm, compact nucleus, pigment and central chromatin. These structures have rounded or pointed ends and are about one and half-time larger than the RBC. 3 The blood film shows the presence of RBC’s, neutrophils, lymphocytes and monocytes. In addition, some of the enlarged RBC’s show within it, large amoeboid coarse haemazoin pigments occupying the entire RBC. 12 to 24 darkly stained merozoites are found within the RBC’s. 4 The Leishman stained thick blood smear shows the presence of coiled structure with a sheath and the absence of nuclei in the tail end. In addition, RBC’s, neutrophils, lymphocytes and monocytes are also seen.

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BOX 68-1 THE PARASITES FOUND IN THE PERIPHERAL BLOOD SMEAR In RBC’s

Plasmodium vivax. Plasmodium falciparum. Plasmodium malariae. Plasmodium ovale. Babesia species.

In leucocytes

Amastigote forms of Leishmania species. Amastigote forms of Trypanosoma species.

In plasma

Microfilaria of Wuchereria bancrofti. Microfilaria of Brugia malayi. Microfilaria of Brugia timori. Microfilaria of Loa loa. Microfilaria of Mansonella perstans.

FIGURE 68-2 The blood film shows the presence gametocyte of Plasmodium falciparum, x1000.

RESULTS AND INTERPRETATION 1 The blood film shows the presence of ring forms of Plasmodium falciparum (Fig. 68-1). 2 The blood film shows the presence of gametocyte of Plasmodium falciparum (Fig. 68-2). 3 Erythrocytic schizonts of Plasmodium vivax in the blood films (Fig. 68-3). 4 Leishman stained thick blood smear with microfilaria of Wuchereria bancrofti (Fig. 68-4).

FIGURE 68-1 The blood film shows the presence of ring forms of Plasmodium falciparum, x1000.

FIGURE 68-3 Erythrocytic schizont stage of Plasmodium vivax in the blood films, x1000.

FIGURE 68-4 Leishman stained thick blood smear with microfilaria of Wuchereria bancrofti, x1000.

KEY FACTS 1 Always thin blood films should be fixed by alcohol or other fixatives before staining. 2 The stains used for staining blood films are of two types: i) One type of stain which has both fixatives and staining reagents e.g., Wright’s, Leishman stains, and ii) Other type of stain which has only staining reagents e.g. Giemsa, Field’s, Jaswant Singh Bhattacharjee stain. Hence with the use of these stains, the thin films should be fixed before staining. 3 Thick blood films should not be fixed because fixation with methanol prevents lysis of RBC’s and dehaemoglobinisation. 4 Thick blood films helps in detection of microfilaria and malarial parasites. It does not serve the purpose of species identification. 5 Thin blood films allows identification of malarial parasite to the species level.

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VIVA 1 Name different Romanowsky’s stains. Ans. Different Romanowsky’s stains are Leishman’s stain, Wright’s stain, Jenner’s stain and Giemsa stain. 2 Name the parasites that can be found in the peripheral blood smear. 3 Name two intra-erythrocytic parasites. Ans. Plasmodium and Babesia. 4 What are the clinical conditions in which examination of a peripheral blood smear may be helpful? Ans. Clinical conditions in which examination of peripheral blood may be useful include: Protozoal infections Malaria. Babesiosis. Chagas disease. African trypanosomiasis. Leishmaniasis. Helminthic infections Lymphatic filariasis. Loiasis. 5 What are differences between thick and thin smears? Ans. Thick smear: It allows examination of larger amount of blood. It is several layers thick. Smear prepared with 2-3 small drops of fresh blood spread over 2 cm diameters. Thin smear:

Allows examination of small amount of blood. Made up of monolayer of RBCs. Smear prepared with a small drop of blood spread along the edge.

FURTHER READINGS 1 Garcia LS. Diagnostic Medical Parasitology. ASM press, Washington D.C. 4th Edition. 2003. 2 Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3rd Edition. 2006. 3 Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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LESSON

69

Concentration of Stool for Parasites

LEARNING OBJECTIVES After completing this practical you will be able to perform: 1 Saturated salt solution flotation method for concentration of stool for parasitic ova and cysts. 2 Formalin-ether sedimentation method for concentration of stool for parasitic ova and cysts.

INTRODUCTION Concentration of stool is recommended when direct examination of stool wet mounts fail to demonstrate any parasites particularly when the number of parasites in stool specimens are low. By concentration methods the helminthic ova and larvae, and protozoal cysts can be concentrated. However, trophozoites cannot be concentrated by any of the concentration methods as they get killed. Various concentration methods are available. Broadly, they can be classified into two groups: flotation techniques and sedimentation techniques. These two approaches are based on the principle of separating parasite from faecal debris and other materials by their differences in the specific gravity. In this chapter, the two commonly used concentration methods, i) Lane’s saturated salt solution floatation method and ii) formalin-ether sedimentation method will be described. Other floatation methods for stool concentration are listed in the table 69-1.

PRINCIPLE In the standard salt solution flotation method, saturated salt solution, a liquid with high specific gravity is used. This helps in separation of protozoal cysts and helminthic eggs from faecal debris. In this method the eggs and cysts of many parasites float because specific gravity of these eggs and cysts are less than specific gravity of the saturated salt solution. The faecal debris are found at the bottom of the container.

The principle of formalin ether sedimentation method is based on recovering helminthic eggs and protozoal cysts in the sediment of the faeces using centrifugation. In this method, the stool sample to be tested is mixed with formalin ether in a test tube and centrifuged. The eggs and cysts, because of the centrifugal force settle down at the bottom of the centrifuge tube and form sediment. The concentrated parasites can be demonstrated in the wet mount preparation of the sediment. The advantages and disadvantages of both the salt solution flotation method and formalin ether sedimentation methods are summarized in the box 69-1.

REQUIREMENTS I Equipments Microscope, centrifuge and discarding jar.

II Reagents and lab wares Microscope slides, broad cover slips, 30 ml glass vials, Pasteur pipette, wooden spatula for both salt solution flotation and formalin-ether sedimentation procedure. In addition to the above, for formalin-ether method alone, centrifuge tubes, glass funnel, measuring cylinder, applicator sticks, surgical gauze and stopper are required. Distilled water, sodium chloride powder (for salt solution method), 5%-10% formalin, ether, saline (for formalin-ether method) Preparation of saturated sodium chloride solution: Saturated sodium chloride solution is prepared by dissolving a spoonful of sodium chloride in 100 ml of distilled water. The salt is dissolved in water using a spatula or a shaker. After all the salt has dissolved completely without a trace, more salt is added till it remains undissolved. This makes saturated sodium chloride solution. III Specimen Stool.

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PROCEDURE Saturated salt solution flotation method 1 Take a walnut sized stool (approx. 1 gram) in a 30 ml glass vial. 2 Add a few ml of saturated sodium chloride solution to the stool and mix it with a broomstick. 3 When the stool suspension is smooth, add a few more ml of salt solution and mix. Note: The steps 2 and 3 are repeated, until the container is nearly full with continuous stirring. 4 Remove any coarse material found floating by a broomstick 5 Add more salt solution little by little using a Pasteur pipette, until a convex meniscus is formed at the top of the container. 6 Then place a broad cover slip over the glass vial, so that it is in contact with the top of the meniscus. 7 Allow it to stand for 20 minutes to 30 minutes. 8 Lift the cover slip from the meniscus with a steady but rapid movement and place on a clean microscope slide. 9 Make a wet mount preparation with one or two drops adherent to the cover slip. 10 Examine the wet mount preparation first under low power objective (10x) of the microscope for parasite eggs. Then focus under the high power objective (40x) to study the morphology and identification of the egg.

Formalin –ether sedimentation method 1 Take a half teaspoon of stool in a 15ml centrifuge tube containing 10 ml of 10% formalin, and allow it to stand for 30 minutes. 2 Filter the faecal suspension through two layers of gauze in a funnel into a 15 ml centrifuge tube. Add saline to the tube to bring the fluid level within several millimeters of the rim of the tube. 3 Centrifuge the tube at 500 g for 10 min. 4 Discard the supernatant. Suspend the sediment in saline, nearly filling up to the brim of the tube. 5 Centrifuge the tube again for 10 min at 500 g. 6 Resuspend the sediment in 7 ml of 10% formalin, and 3 ml of ether. 7 Close the tube with a stopper and shake it well for 30 seconds. Note: Hold the tube in such a way that the stopper is held away from the face. 8 Remove the stopper carefully. 9 Centrifuge the tube at 500 g for 10 minutes. Allow the tube to stand 5 min. Note: After centrifugation, four layers are formed namely, i) first sediment layer at the bottom of the tube containing parasitic cysts or eggs, ii) second layer of formol saline, iii) third layer of faecal debris on the top of the formol saline layer and iv) lastly a top layer of ether. 10 Remove the plug of faecal debris by piercing all around with an applicator stick, taking care not to disrupt the debris.

11 Discard all the fluid into the discarding jar by one firm swing, leaving behind one or two drops of fluid with the sediment. 12 Mix the sediment with the fluid using an applicator stick. Then with a Pasteur pipette, aspirate a little of the sediment fluid and examine by making the saline and iodine wet mount preparations. 13 Examine the wet mount preparation first under low power objective (10 x) of the microscope for parasite eggs and cysts. Then focus under the high power objective (40x) to study the morphology and identification of the egg and cysts.

QUALITY CONTROL Wet mounts are prepared from a stool specimen known to be positive for ova and cysts and compare with the wet mounts obtained after the formol ether sedimentation technique for similar morphology of eggs and cysts.

OBSERVATIONS 1 A 10-15µm sized spherical structure seen possessing 1-4 nuclei. 2 A 15-30µm sized spherical structure seen possessing 1-8 nuclei. 3 A 6-10µm size oval ellipsoidal shaped structure seen with 4 nuclei remains of axoneme and flagella. 4 A 60µm × 40µm size oval shaped, non bile stained structure seen with a thin transparent hyaline shell membrane enclosing 7-8 blastomeres, and clear space between the shell membrane and blastomeres. 5 A 60-75µm × 40-50µm size rounded, bile stained structure seen with a thick translucent shell with an albuminous coat containing a large conspicuous unsegmented ovum with a clear space at each pole.

RESULTS AND INTERPRETATION 1. 2. 3. 4. 5.

Cyst of Entamoeba histolytica /dispar. Cyst of Entamoeba coli. Cyst of Giardia intestinalis. Egg of hook worm. Fertilized egg of Ascaris lumbricoides.

Table 69-1 Other floatation methods for concentration of stool Zinc sulphate floatation method. Magnesium sulphate floatation method. Sheather’s sucrose floatation method.

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BOX 69-1 ADVANTAGES AND DISADVANTAGES OF THE CONCENTRATION METHODS Saturated salt floatation method Advantages Simple procedure. Inexpensive method and easy to perform. Good method for concentration of parasitic eggs. Can be done at any level of the laboratory. Least subjective to technical error. Disadvantages Not suitable for demonstration of unfertilized eggs of Ascaris, operculated eggs of trematodes, larvae of Strongyloides and eggs of Taenia species. High specific gravity of the fluid may cause distortion of the morphology of the parasitic eggs and cysts. Formalin-ether sedimentation method Advantages More sensitive method. Good method for both cysts and ova. Morphology of eggs and cysts preserved. Disadvantages Technically cumbersome. Ether is highly inflammable and not easily available.

KEY FACTS 1 Concentration of stool is the method of choice, when the number of parasites in the stool specimens are less and cannot be demonstrated in stool wet mount preparation. 2 Trophozoites of protozoa cannot be concentrated by any of the concentration methods. 3 Stool concentration by salt floatation and formol ether sedimentation are the two reliable methods of concentration of parasites in stool. It increases sensitivity of stool microscopy. 4 Salt floatation is a better method for concentration of helminthic eggs. 5 In formol ether procedure, formol saline is used to fix the cysts of protozoa and eggs, thus unaltering morphology of these structures. These also kill the parasitic eggs and cysts. Ether is used to dissolve and extract the fat and other debris in the stool.

VIVA 1 2 3 4 5 6

What are the advantages of floatation method? What are the disadvantages of floatation method? What are the advantages of sedimentation method? What are the disadvantages of sedimentation method? Name the other floatation methods that can be employed as stool concentration method. Which is the stool concentration method employed for recovery of coccidian parasites? Ans. The stool concentration method employed for the recovery of coccidian parasites is the Sheather’s sucrose floatation method.

FURTHER READINGS 1 Garcia LS. Diagnostic Medical Parasitology. ASM press, Washington D.C. 4th Edition. 2003. 2 Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3rd Edition. 2006. 3 Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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70

Culture of Stool for Entamoeba histolytica

LEARNING OBJECTIVES

REQUIREMENTS

After completing this practical you will be able to:

I Equipments Bacteriological incubator and inspissator.

1 Culture stool for Entamoeba histolytica.

INTRODUCTION For the cultivation of Entamoeba histolytica various media have been used. The media have been classified broadly into two groups: polyxenic (polybacterial culture) and axenic (bacteria –free culture) medium. The polyxenic media are Boeck and Drbohlav’s Locke-egg-serum (LES) medium, Balamuth’s, Nelsons’ or Robinson’s medium. Boeck and Drbohlav’s Lockeegg serum (LES) medium is a polyxenic medium commonly used for culture and isolation of the amoebae from the stool specimens for diagnostic purposes. Trophozoites of E. histolytica usually appear in large numbers within 48 hours of inoculation.

PRINCIPLE Polyxenic culture medium is usually composed of egg and Locks’ solution mixture supplemented with serum, starch and bacterial flora, which provide nourishments for the growing parasites. This is used routinely for diagnosis of intestinal amoebiasis by isolation of the amoeba from the stool. Axenic culture is a bacteria-free culture medium. This is used to study i) the pathogenicity of amoebae, ii) testing antiamoebic drugs in vitro, iii) immunological properties of amoebic antigen and iv) prepare axenic amoebic antigen for use in the immunodiagnosis of amoebiasis. Axenic culture is not used for diagnosis by isolation of amoebae from the stool. In this chapter polyxenic culture of E.hsistolytica in Boeck and Drbohlav’s medium will be described.

II Reagents and lab wares Boeck and Drbohlav’s Locke-egg-serum (LES) medium, heat inactivated bovine serum, screw cap tubes, and antibiotics solution. Preparation of polyxenic Boeck and Drbohlav’s medium: 1 Break four eggs into a sterile flask containing glass beads, after that they are washed and shells are wiped dry with 70% alcohol. 2 Then add 50 ml of Locke’s solution and shake the mixture until homogenous. 3 Dispense the medium in tubes, such that a slant of 1 to 1.5 inches are produced at the bottom of the tube. 4 Plug the tubes and place the tubes in a slant position in an inspissator at 70°C until the slant solidifies. 5 Autoclave the tubes at 15 lb pressure for 20 minutes. 6 Prepare a mixture of 8 parts of sterile Locke’s solution with 1 part sterile inactivated bovine serum. 7 Sterilise the mixture by filtration and incubate at 37°C for 24 to 48 hours as a sterility check before use. 8 Cover the slants of the medium to a depth of 1 cm with 6 ml to 8 ml of the sterile lock’s solutions. 9 Add a loopful of sterile rice powder/starch to each tube along with loopful of Escherichia coli colony. 10 Add antibiotics solution (penicillin, 1000 units/ml, streptomycin 2 mg/ml and acriflavine, 0.1 ml of 0.02%) to the medium to inhibit the overgrowth of commensal bacteria present in the stool. III Specimen Stool in cases of suspected amoebic dysentery, and pus in cases of amoebic liver abscess.

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PROCEDURE

QUALITY CONTROL

1 Inoculate a loopful of the stool sample or liver pus on to the slant of the two medium tubes. 2 Incubate the tubes in an incubator at 37°C for 24 hours to 48 hours. 3 Observe the cultures regularly at 2, 3 and 4 days by examining 0.1 ml of sediment under light microscope for characteristic motility of the trophozoites. Note: Although the initial culture may appear negative, subcultures may reveal amoebae. 4 Examine the growth of amoebae by a wet mount preparation of culture fluid.

A known positive culture of E. histolytica already maintained by serial sub cultivation in the culture medium.

OBSERVATIONS Presence of 8–30 µm structure, actively motile with hyaline, finger-shaped pseudopodia, clearly differentiated cytoplasm into ectoplasm and endoplasm, and cytoplasmic inclusions such as red blood cells, leucocytes and tissue debris.

RESULTS AND INTERPRETATION Trophozoites of E. histolytica is grown in the culture.

KEY FACTS 1 2 3 4 5

Cultivation of E. histolytica can be done using both polyxenic and axenic culture medium. In polyxenic culture medium (polybacterial culture) bacterial flora serve as rich source of nutrients for the feeding amoebae. In axenic culture medium (bacteria-free culture), addition of special vitamin mix provides a rich source of nutrition. Axenic culture medium is not used routinely for culture and isolation of the amoebae from stools specimens. Axenic culture medium is employed to study pathogenicity of amoeba, in vitro testing of efficacy of anti-amoebic drugs and preparation of axenic amoebic antigen for immunodiagnosis of amoebiasis.

VIVA 1 2 3 4

Name the axenic culture medium used for culture of E. histolytica. Define axenic and polyxenic culture medium. What are the uses of axenic culture of E. histolytica? List the polyxenic culture media and the uses of polyxenic culture of E. histolytica.

FURTHER READINGS 1 Garcia LS. Diagnostic Medical Parasitology. ASM press, Washington D.C. 4th Edition. 2003. 2 Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3rd Edition. 2006. 3 Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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UNIT

X Mycology Introduction Lesson 71 Cultivation of Fungi Lesson 72 Gram’s Staining for Fungi Lesson 73 Lactophenol Cotton Blue (LPCB) Wet Mount Lesson 74 Potassium Hydroxide Wet Mount Lesson 75 India Ink Preparation Lesson 76 Slide Culture Lesson 77 Germ Tube Test Lesson 78 Urease Test Lesson 79 Carbohydrate Assimilation Test Lesson 80 Carbohydrate Fermentation Test Lesson 81 Identification of Common Fungi

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Introduction The fungi are now recognized as significant causes of morbidity and mortality. They have emerged as important etiological agents of opportunistic infections and full-fledged diseases. Today, the incidence of the fungal infections has increased enormously, due to underlying predisposing factors such as immunocompromised situations. The prognosis among these patients is very poor and therefore an early diagnosis and treatment is essential. The symbiotic relation of the fungi, like other organisms, can be divided into three modes of their existence namely mutualism, commensalisms and parasitism. The aged patients, whose life span has been extended by treatment of cancer or other debilitating diseases, are more susceptible to secondary fungal infections than the young individuals. In addition many factors directly or indirectly decrease the accuracy of data on fatal mycoses cases. The fungal infections are not usually transmitted sexually like those of the viral, bacterial or parasitic diseases. Till date there is no ideal vaccine available to control any fungal infections. Simultaneously, the reported deaths from the fungal infections have maintained an approximately constant numerical ratio to the general increase in the population. The nosocomial fungal infections have been recognized as a significant ground for adverse patient outcome and a major public health problem. Nosocomial blood stream infections are particularly serious and there is evidence to suggest that these infections are becoming more common. Nosocomial mycoses have developed especially in association with or as a consequence of the extraordinary progress in the management of seriously ill patients. However, despite this increase, there has been comparatively little progress in understanding the pathogenesis of nosocomial fungal infections or in their prevention, diagnosis and treatment. The fungal diseases can be classified according to the primary site of infection as follows: 1 Superficial mycoses: The infection is limited to the outer most layers of the skin and its appendages. The immune response is rarely induced. 2 Cutaneous mycoses: The infection extends deeper into the epidermis and it also invades hair and nails. It evokes a high inflammatory response in the host. 3 Subcutaneous mycoses: The infection is due to the pathogenic organism of low virulence and usually following traumatic injury. It involves the dermis, subcutaneous tissues, muscles and fasciae. 4 Systemic mycoses: The infection originates primarily at one site and disseminate systemically to other body sites. 5 Besides these a fifth group opportunistic has come into focus because of increasing use of immunosuppressive therapy or AIDS epidemic. These infectious agents are of low pathogenic potential and produce disease only under unusual circumstances, mostly involving host debilitation. Fungal infections can be diagnosed by their demonstration, isolation and final identification from clinical specimens. Diagnosis of fungal infections is made by direct and indirect methods. Direct methods include the demonstration of fungi or their components in body tissues or fluids. Wet mount of clinical specimens is a very useful method for demonstration of fungi in clinical specimens. Various wet mount preparation used in a mycology laboratory include lactophenol cotton blue (LPCB) wet mount, potassium hydroxide (KOH) wet mount and India ink preparation.

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71

Cultivation of Fungi

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Select appropriate medium to grow fungi from clinical specimens. 2 Isolate fungi from clinical specimens in the laboratory.

INTRODUCTION Different kinds of media are available to grow and isolate different types of fungi. List of different media used for culture of fungi are summarized in the table 71-1. Sabouraud’s dextrose agar (SDA) (Fig. 71-1) is the most frequently used media in a diagnostic mycology laboratory (Box 71-1).

PRINCIPLE Clinical specimens should be processed promptly and plated on to isolation media as a means to recover fungi that may be causing disease. Appropriate media and incubation temperatures are selected to allow for the growth of pathogenic and opportunistic yeasts and fungi.

REQUIREMENTS

III Specimen Skin and nail scrapings, hair, and exudate from lesions, urine, sputum, bronchial washings, biopsied materials and CSF.

PROCEDURE 1 Choose and take two appropriate sterile medium tubes and label RT (room temperature) and 37°C. 2 Take processed specimen and inoculate a loop full of specimen in each tube. 3 Incubate RT tube at room temperature and 37°C tubes at 37°C temperature.

QUALITY CONTROL Prepared media should be incubated and then used to ensure sterility. Quality of sterilization should be checked periodically and appropriate pH should be adjusted for best results. The control strains should be used to grow and see if growth is perfect in prepared medium.

OBSERVATIONS Observe both the test tubes and control tubes everyday for upto 30 days.

I Equipments Water bath, autoclave/hot air oven for sterilization, pH meter or pH paper.

1 Observe for fungal colonies. 2 Tubes growing bacterial colonies are discarded.

II Reagents and lab wares Glassware for preparation of media, tubes for aliquots of media, Bunsen burner, inoculating loop, glass marking pencil, appropriate sterile media (Table 71-1).

RESULTS AND INTERPRETATION Growth of fungal colonies in the tube inoculated with test specimen indicates the patient is infected with fungus (Fig. 71-1).

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Table 71-1 List of media used for fungal culture Sabouraud’s dextrose agar (SDA) Sabouraud’s dextrose agar with antibiotics Ascospore medium Bennett’s agar for Nocardia Blood agar base Casein agar base for identification of Trichophyton species Casein agar for Nocardia Casitone medium of Howell and Pine for Actinomyces Carbohydrate broth for Candida Czapek – Dox medium Gelatin medium for Nocardia Malt – Yeast extract agar (Wicker ham) Medium 199 for Nocardia munutissima Mohapatra and Pine medium for Nocardia Potato glucose agar Rice agar with Tween 80 Rice medium for Microsporum species Salvin’s YP medium for yeast phase of Histoplasma capsulatum Tarshis’ penicillin blood Agar Trypticase soy agar Tyrosine agar Xanthine Agar

BOX 71-1 SABOURAUD’S DEXTROSE AGAR Sabouraud’s dextrose agar It consists of 40 gm of glucose / dextrose, 10 gm of neopeptone and 35 gm of agar in 1000 ml of distilled water. pH is 5.5 – 6.0. Sabouraud’s dextrose agar with antibiotics It consists of 20 gm of glucose/dextrose, 10 gm of neopeptone and 20 gm of agar in 1000 ml of distilled water. pH is 7.0. Sterilised by autoclaving. It contains 40 mg of chloramphenicol and 40 mg of cycloheximide (actidione).

FIGURE 71-1 SDA with Candida albicans colonies.

KEY FACTS 1 2 3 4

Choose appropriate medium based upon clinical history of the patient. Always inoculate in duplicate tubes and incubate one at room temperature and another at 37°C. Media should be sterile. pH should be appropriate.

VIVA 1 What are the different types of media used in fungal culture? 2 What are the most commonly used media in laboratory for routine diagnosis of fungal infections? Ans. Sabouraud’s dextrose agar (SDA) is the most frequently used media in a diagnostic mycology laboratory The other commonly used media include carbohydrate broth, Czapek- Dox medium, gelatin medium, potato glucose agar, trypticase soy agar and urea agar. 3 How do you sterilize different media used for culturing fungi? Ans. Most media used for fungal culture are sterilized by autoclaving at 15 lbs for 15 mins. 4 How do you test for the proper working of different media? Ans. The media that are prepared can be tested to see for its proper working by inoculating standard strains of appropriate fungi and incubating them both at 37°C and at room temperature.

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000.

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72

Gram’s Staining for Fungi

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Stain fungal smears by Gram’s staining method.

INTRODUCTION Gram stain was devised by Christian Gram in 1884 as a method of staining bacteria in tissues. Fungal material appears Gram positive.

PRINCIPLE The Gram reaction is dependent on the permeability of the fungal cell wall, to the dye-iodine complex like those of bacteria (refer chapter 7).

REQUIREMENTS I Equipments Compound light microscope. II Reagents and lab wares Bunsen flame, loop wire, methyl violet (basic dye), Gram’s iodine (mordant), 95% ethanol (decolourising agent), and 1% safranine or dilute carbol fuchsin (counter stain).

Sabouraud’s dextrose agar and place on the clean slide with a bacteriological loop. 3 Then with circular movement of the loop, spread the cell suspension into a thin area. 4 Allow the smear to air dry. 5 Heat fix the smear while holding the slide at one end, and quickly passing the smear over the flame of Bunsen burner two to three times.

II Staining Procedure 1 Heat fixes the smear by passing the slide 2–3 times gently over the flame with the smear side up. 2 Cover the smear with the methyl violet. Allow it to stand for one minute. 3 Rinse the smear gently under tap water. 4 Cover the smear with Gram’s iodine and allow it to stand for one minute. 5 Rinse the smear again gently under tap water. 6 Decolourise the smear with 95% alcohol for 15 to 20 seconds. 7 Rinse the smear again gently under tap water. 8 Cover the smear with dilute carbol fuchsin for 30 seconds to 1 minute. 9 Rinse the smear again gently under tap water and air dry it. 10 Observe the smear first under low power (10x) objective, and then under oil immersion (100x) objective. 11 Record the observations in the note book.

QUALITY CONTROL III Specimen Candida albicans culture on Sabouraud’s dextrose agar.

PROCEDURE I Preparation of fungal smear: 1 Take clean, and grease free glass slides for making the smears. 2 Take one or two loopful of C. albicans culture on

On the same slide, at one end a thin control smear of mixture of Staphylococcus aureus (Gram positive bacteria) and Escherichia coli (Gram negative bacteria) is made and at other end of the slide test smear is made. The slide with control and test smears is stained by Gram’s staining. The appearance of purple coloured Gram positive bacteria and pink coloured Gram negative bacteria in the control smear indicates proper staining technique and stained test smear is compared with it.

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OBSERVATION Presence of Gram positive budding yeast cells.

RESULTS AND INTERPRETATION The stained smear contains Gram positive yeast , C. albicans (Fig. 72-1). FIGURE 72-1 Gram stained smear of Candida albicans, x 1000.

KEY FACTS 1 Gram staining is a differential stain, which can also be used for detection of fungi in clinical specimens. 2 Gram staining gives preliminary indication of infection. 3 Tissue cells, leucocytes and the debris of inflammatory exudates all stain pink in Gram’s stained smears.

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000.

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73

Lactophenol Cotton Blue (LPCB) Wet Mount

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Prepare lactophenol cotton blue (LPCB) wet mount of fungal colony. 2 Demonstrate fungi and fungal elements under the microscope in LPCB wet mount preparation.

INTRODUCTION Fungal infections can be diagnosed by their demonstration in clinical specimens. LPCB staining wet mount is the most commonly used method adopted in a mycology laboratory to identify filamentous fungi.

PRINCIPLE Identification of filamentous fungi is made by their characteristic microscopic morphology such as shape, size, arrangement of spores and hyphae. There are three different preparations of LPCB mounts as mentioned below: 1 Scotch tape preparation. 2. Tease mount preparation, and 3 Slide culture preparation. In case of scotch tape preparation, the fungal elements are undisturbed whereas in tease mount preparation, the fungal elements are disturbed but finer details of the fungus can be visualized. Slide culture (Ref. chapter 75) is the most advantageous in that, the fungus as it grows on the medium can be visualized. In this chapter, LPCB wet mount of both Scotch tape preparation and tease mount preparations will be described.

REQUIREMENTS I Equipments Microscope.

II Reagents and lab wares Glass Petri dishes, slide, cover slip, straight/bent wire and needle, lactophenol cotton blue (LPCB) stain. Preparation of Lactophenol cotton blue stain: Weigh and add 20 gm of phenol crystals, 20 ml of lactic acid, and 40 ml of glycerol to 20 ml of distilled water. Dissolve the ingredients by heating the container in a hot water bath. Add 0.05 gram cotton blue. III Specimen Rhizopous culture on Sabouraud’s dextrose agar.

PROCEDURE Scotch tape preparation 1 On a clean glass slide, place one drop of LPCB. 2 Touch the adhesive side of the tape of transparent scotch tapes on the surface of the colony at a point intermediate between its centre and periphery. 3 Fix the adhesive side of the tape over an area on the glass slide containing the LPCB. 4 Examine the preparation under 10x and 40x of a light microscope.

Tease mount preparation 1 Place a drop of LPCB on a clean glass slide. 2. Remove a small portion of the colony and the supporting agar at a point between the centre and periphery and place it in the drop of LPCB. 3 With a needle, tease the fungal culture first and spread in the LPCB. 4 Examine microscopically after giving sufficient time for the structures to take up the stain, usually 30 mins.

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Lactophenol Cotton Blue (LPCB) Wet Mount

QUALITY CONTROL Preparation of lactophenol cotton blue should be done by heating the ingredients in a waterbath.

types of morphological structures including hyphae and spores. This should be thoroughly differentiated and the fungus identified. Fast growing fungi in case of slide culture preparation will give satisfactory results in 24–48 hr.

OBSERVATIONS 1 The stained preparation should be observed under 10x or 40x, for presence of mould. 2 Fungi appear as dark blue stained mycelium (Fig. 73-1).

RESULTS AND INTERPRETATIONS The fungal element grown should be observed and results interpreted depending on the morphology of the hyphae and the spores. Different fungi under LPCB wet mount will show different

FIGURE 73-1 LPCB mount of fungi, x 400.

KEY FACTS 1 The fungal culture should be first is teased well and then spread in LPCB for better results.

VIVA 1 What is lactophenol cotton blue and how is it prepared? 2 What are three different methods of wet mounts used and their advantages in a lactophenol cotton blue preparation? 3 What are the functions of each component of lactophenol cotton blue stain? Ans. a Lactic acid: Helps in preserving the morphology of the fungal element. b Phenol: Acts as disinfectant. c Cotton blue: Stains the fungal elements. d Glycerol: Hygroscopic agent. It prevents drying. 4 What is the other use of LPCB apart from examination of fungi? Ans. LPCB is also used in wet mount of stool for intestinal parasites.

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000.

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74

Potassium Hydroxide Wet Mount

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to:

1 Emulsify the specimen in a drop of 10% KOH on a microscopic slide with the help of a loop. 2 Apply gentle heat by passing the slide over a Bunsen flame for 3–4 times. 3 Cover the smear with the cover slip. 4 Leave it for 5–10 min. 5 Examine the slide under low (10x) and high power (40x) magnifications 6 Examine the slide for 15–20 min. for demonstration of shining fungal elements.

1 Prepare potassium hydroxide (KOH) wet mount of clinical specimens. 2 Demonstrate the presence of fungal elements in the given clinical specimen by KOH wet mount preparation.

INTRODUCTION The potassium hydroxide (KOH) wet mount preparation is very useful for the presumptive diagnosis of the type of fungal infection. The procedure also helps in the selection of appropriate culture media for the isolation of etiological fungal agent.

PRINCIPLE The KOH clears out the background scales or cell membranes that may be confused with fungal hyphal elements in microscopy of clinical specimens. Gentle heating also accelerates clearing of artifacts.

QUALITY CONTROL 1 10% KOH should be prepared at the right concentration. 2 Emulsification of specimen should be homogenous in KOH solution.

OBSERVATIONS Shining fungal elements shall be observed in microscopy of the clinical specimens.

REQUIREMENTS RESULTS AND INTERPRETATIONS I Equipments Microscope. II Reagents and lab wares Glass Petri dishes, slide, cover slip, straight/bent wire, needle, Bunsen flame and 10% KOH. III Specimen Pus from draining sinuses, aspirate from nasal sinuses, respiratory specimen, skin scrapings, nail scrapings, hair, corneal scrapings, material from external ear, etc.

Different fungi will have different morphological forms (yeasts, cells with pseudo hyphae, budding, septate, and aseptate hyphae, granules, etc.) which can be clearly seen in a KOH wet mount. Interpretation of results should be done by critical analysis of the type, size and color of fungal elements which will be different for different fungi.

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KEY FACTS 1 The KOH clears out the background scales or cell membranes that may be confused with fungal hyphal elements in microscopy of clinical specimens. 2 The KOH should be prepared at the right concentration (10%). 3 Emulsification of specimen should be homogenous.

VIVA 1 What percentage of KOH should be used? 2 What is the function of KOH? 3 How do you interpret results of a KOH wet mount?

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000. pp. 232.

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75

India Ink Preparation

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to:

1 2 3 4

1 Prepare India ink wet mount of cerebrospinal fluid (CSF). 2 Demonstrate the presence of capsulated yeast, Cryptococcus neoformans in the CSF.

INTRODUCTION Capsule is a protective layer found around some bacteria and some fungi like C. neoformans. Hence demonstration of capsule by India ink preparation, especially in an emergency conditions, in cerebrospinal fluid (CSF) is a very useful procedure for diagnosis of meningitis caused by C. neoformans. An early diagnosis will help for prompt treatment of the condition.

PRINCIPLE India ink is used as a negative stain preparation. When used in wet mount preparation of the CSF, the background appear black, and the unstained capsule of C. neoformans appears as a white halo around the yeast cells in microscopy.

REQUIREMENTS

Put a drop of CSF on the microscopic slide. Put a drop of India ink to the CSF on the microscopic slide. Emulsify the specimen with India ink on the slide. Place a cover slip over the preparation, taking care not to trap air bubbles in the preparation. 5 Blot dry the excess fluid. 6 Examine the slide under low (10x) and high power (40x) magnifications

QUALITY CONTROL 1 India ink preparation in distilled water should be made exactly to 0.5%. 2 Care should be taken not to trap air bubbles which will mimic capsules of yeast cells.

OBSERVATIONS The India ink preparation is observed under microscope and noted for presence of clear halo around yeast cells (Fig. 75-1).

RESULTS AND INTERPRETATION Since India ink stains the background and leaves a clear halo around the cells, preparations with such appearance can be confirmed to have yeast cells with capsules and the organism may be identified as C. neoformans.

I Equipments Microscope. II Reagents and lab wares Microscopic slide, cover slip, glassware, loop wire and 0.5% India ink in distilled water. III Specimen Cerebrospinal fluid (CSF).

FIGURE 75-1 India ink preparation showing Cryptococcus neoformans with capsule, x 400.

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Indian Ink Wet Mount Preparation

KEY FACTS A clear distinction should be made between capsules and air bubbles, trapped between the cover slip and slide.

VIVA 1 What is the percentage of India ink used? 2 What is the advantage of India ink preparation and how is it useful in emergency mycology laboratory? Ans. India ink preparation is one of the best methods to demonstrate the presence of capsule in case of capsulated yeasts. The demonstration of capsulated yeast like C. neoformans in CSF is significant in emergency mycology laboratory for prompt administration of antifungal agents. 3 How to interpret results of India ink preparation? Ans. The India ink preparation should be critically interpreted for the presence of a clear capsule around the yeast cell. This should not be confused with any air bubble that might be present.

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000.

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Slide Culture

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Perform slide culture of the fungal preparation. 2 Mount the cover slip after the growth of fungus. 3 Demonstrate fungal morphology without disturbing the aerial hyphae and conidiophores, if present.

INTRODUCTION Slide culture is a very useful technique to study undisturbed morphological details of fungi, particularly relationship between reproductive structures like conidia. An entire fungal colony can be demonstrated within a short period of time with the use of minimum materials.

PRINCIPLE Slide culture is a very useful technique in identification of the type of fungi. The fungal element that is to be identified will produce characteristic hyphae and spores, when incubated on a suitable growth medium. This can be visualized undisturbed using this technique. The mould that is to be cultured is inoculated onto a small piece of an agar below a cover slip. The whole setup is kept in a Petri dish with moisture. The cover slip after incubation is lifted, stained and observed under a microscope for identification of the fungi.

REQUIREMENTS I Equipments 1 Slide culture set (Fig. 76-1). (a) Into a Petri dish, place one piece of filter paper slightly smaller in diameter than the Petri dish. (b) Place a V-shaped glass rod on the filter paper. (c) Place a 1 by 3 inch glass slide and a 22 mm square, No. 1 glass cover slip on top of the filter paper.

(d) Wrap the preparation in Kraft 20 paper for sterilizing in hot air oven. Several of these setups should be kept ready on hand. 2 Sterile test tube with rimless mouth and an inside diameter of approximately 15 mm. 3 Standard laboratory glassware. II Reagents Sterile distilled water and Petri dish containing Sabouraud’s agar (or other medium of choice) to a depth of 2 mm. III Specimen Fragments of mould to be cultured is used as the specimen.

PROCEDURE 1 From the Petri dish containing Sabouraud’s agar cut out one square cm block of agar for each slide culture to be inoculated. 2 With the flat side of a sterile bacteriological loop, or with a spatula, place an agar block in the centre of the slide in the slide culture set up. 3 With a probe, inoculate around the periphery of the agar block, three to four fragments of the mold to be cultured. 4 With forceps, the tips of which have been flamed, place the cover slip on the agar block. 5 With a pipette, thoroughly moisten, but not to excess, the filter paper with sterile distilled water. 6 Incubate the slide culture at room temperature. 7 Remove the slide culture from the Petri dish and dry the bottom of the slide with a tissue. 8 When growth appears peneath the cover slip. Take a slide place a drop of LPCB, place the cover slip removed from the block on the LPCB. 9 Place the slide on the microscope stage and examine. The aerial hyphae including the conidiophores will be seen to grow along the undersurface of the cover slip

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QUALITY CONTROL

RESULTS AND INTERPRETATION

1 All the materials should be sterilized and checked for sterility before use. 2 Distilled water to be used should be checked for sterility . 3 Components of the medium used should be adjusted according to standard procedure.

Small spore bearing fungi make beautiful permanent mounts. Some large spore bearing organisms like Microsporum gypsum do not stain as well. With the type of hyphae, arrangement of conidiophores, staining characters etc., the final interpretation of the fungal type can be made.

OBSERVATIONS Usually a minimum of 48 hr. is needed before a slide culture shows growth of aerial hyphae. Thus, the culture may be examined after 48 hours incubation and as frequently thereafter as necessary.

FIGURE 76-1 Slide culture set.

KEY FACTS 1 The Petri dish chamber should be always moist. 2 Agar used should support growth of the suspected fungus. 3 Sterility should be maintained to the maximum.

VIVA 1 What are the requirements for slide culture? Ans. The requirements for slide culture include Petri dish, piece of filter paper slightly smaller in diameter than the Petri dish, V shaped glass rod, 1 by 3 inch glass slide, 22 mm square, No. 1 glass coverslip; Kraft 20 paper, sterile distilled water, sterile test tube with rimless mouth and an inside diameter of approx. 15 mm, Sabouraud’s agar (or other medium of choice) to a depth of 2 mm and standard laboratory glassware and. 2 When is slide culture used in routine diagnosis of fungal infections? Ans.Slide culture can be used in routine diagnosis of fungal infections when examination of the entire fungal colony is required without disturbing the aerial hyphae and conidiophores, if present. Slide culture is a technique used to study an entire fungal colony within a short period of time with the use of minimum materials. 3 What are the main advantages of slide culture? Ans. Slide culture is a convenient method to demonstrate an entire colony without disturbing the aerial hyphae and conidiophores of fungi such as Aspergillus species, Penicillium species etc.

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000. pp. 232.

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LESSON

77

Germ Tube Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the production of germ tube by Candida species.

INTRODUCTION Candida species are usually found as normal flora of the oral cavity and gastrointestinal tract of man. These may be isolated from respiratory secretions, gastric washings, stool, vaginal secretions, urine, skin, nail, etc. Under normal conditions Candida species are not pathogenic. However, in case of immunocompromised individuals, and certain other situations (Box 77-1), Candida species can cause a wide variety of opportunistic infections. Germ tube test is a very simple and efficient test to distinguish pathogenic Candida from non-pathogenic ones. It is very widely used in diagnostic laboratories because of its reproducibility.

PRINCIPLE Germ tube is an initial hypha from a sprouting conidia, spore or yeast. Formation of germ tube can be demonstrated by inoculating rabbit, fetal calf or human serum with a small quantity of growth of Candida species. The suspension is then incubated at 37°C for a minimum of 1½–2 hours, after which a drop is examined under the microscope for the germ tube. Germ tubes are produced by C. albicans, C.stellatoidea and rarely C. tropicalis. At times, some strains of C. albicans isolated from the patients with antifungal drugs or patients with cancer do not produce germ tubes.

REQUIREMENTS I Equipments Microscope.

II Reagents and glass wares Standard laboratory glassware, and test tubes 12×75 mm, human, foetal calf or rabbit serum. III Specimens 24 hour culture of suspected fungal colony on Sabouraud’s dextrose agar to be tested. 24 hour culture of known strains of C. albicans and C. parapsilosis colony on Sabouraud’s dextrose agar.

PROCEDURE 1 Take three test tubes and label as 1, 2 and 3. 2 Add 0.5ml. of serum to each of the test tube. 3 Take a half of a single colony to be tested by using a sterile loop, and mix with serum in the test tube 1. 4 Similarly, take a half of C. albicans single colony by using a sterile loop, and mix it with serum in the test tube 2. 5 Similarly, take a half of C. parapsilosis single colony by using a sterile loop, and mix it with serum in the test tube 3. 6 Incubate all the tubes at 37°C for a maximum of 1½ hrs. 7 Place one drop of suspension from tube 1, 2, and 3 onto 3 different slides and place cover slips over the drops. 8 Examine the slide under low (10x) and high power (40x) magnifications.

QUALITY CONTROL 1 Positive control for germ tubes: C. albicans. 2 Negative control for germ tubes: C. parapsilosis. 3 Human or rabbit serum should be checked for contamination prior to use. 4 Control organisms should be tested individually for production of germ tubes.

OBSERVATIONS Under the microscope, the whole field under the cover slip is examined for any cell showing production of germ tube (Fig. 77-1).

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Germ tubes are seen as long tube like projections extending from yeast cells. This should be differentiated from pseudohyphae (Table 77-1).

RESULTS AND INTERPRETATION Tube 2 will show production of germ tube and Tube 3 will not. The drop from tube 1 should be read and compared with these controls. Tube 2 contains C. albicans and hence shows germ tube production while tube 3 does not show production of germ tube since it contains C. parapsilosis. Tube 1 should be interpreted with care by observing for the presence or absence of germ tube and should be compared with tube 2 and tube 3.

FIGURE 77-1 Germ tube test, x 400.

BOX 77-1 PREDISPOSING FACTORS FOR CANDIDIASIS

Table 77-1 Differences between germ tubes and pseudohyphae

Immunosuppression Long term antibiotic therapy Use of oral contraceptives Pregnancy Premature birth Obesity Diabetes mellitus Immunocompromised status

Germ tubes

Pseudohyphae

No constriction at the site of attachment.

Constriction at the site of attachment.

Non-septate with parallel sides.

Septate and not necessarily with parallel sides.

KEY FACTS 1 2 3 4

Germ tube is a useful test to identify C. albicans and few other species which are pathogenic. Rabbit, foetal calf or human serum can be used for demonstrating germ tube formation. Germ tubes are produced by C. albicans, C.stellatoidea and rarely C. tropicalis. At times, some strains of C. albicans isolated from the patients with antifungal drugs or patients with cancer do not produce germ tubes. 5 C. tropicalis may show germ tube formation after 3 hours with a constriction at the base of the germ tube.

VIVA 1 What is a germ tube and how is it significant in routine identification of fungi? 2 Why is germ tube test important in a mycology laboratory? 3 What are the advantages of a germ tube test?

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000.

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LESSON

78

Urease Test

LEARNING OBJECTIVES

III Specimen C. neoformans culture.

After completing this practical you will be able to: 1 Demonstrate production of the enzyme urease by the fungus Cryptococcus neoformans.

INTRODUCTION Some fungi produce the enzyme urease that hydrolyses the urea releasing ammonia into the medium. Ammonia in turn produces a change in the pH of the medium that can be detected by the colour change in the indicator dye. This test can be used to differentiate different groups of fungi.

PRINCIPLE Urea is a diamide of carbonic acid. Urease, the enzyme produced by the fungi, hydrolyses urea and releases ammonia and carbon dioxide. Ammonia reacts in solution to form ammonium carbonate, which is alkaline, leading to an increase in the pH. Phenol red that is incorporated in the medium changes its colour from yellow to red in alkaline pH, thus indicating the presence of urease activity.

PROCEDURE 1 Pick up one colony of C. neoformans. 2 Inoculate Christensen’s urea agar slope with these fungal colonies. 3 Incubate the tube at 37°C for 2 days. 4 Observe any change of colour in the inoculated medium.

QUALITY CONTROL Positive control: Trichophyton mentagrophytes (urease positive fungi). Negative control: Candida albicans and Trichophyton rubrum (Urease negative fungi). An un inoculated medium is incubated along with the test to compare the colour change.

OBSERVATION Examine after 2 days of incubation. The test should not be considered negative till after 2 days of incubation. The un inoculated medium is colour less. In a positive test, after incubation, the colour of the medium changes to purple pink.

REQUIREMENTS

RESULTS AND INTERPRETATION

I Equipments Incubator.

Positive reaction is detected within 1 to 2 days of incubation. When positive, medium shows growth of the colony and the color of the test medium as well as positive culture medium changes to purple pink. No colour change is seen in negative culture medium. The test is considered negative if no colour change of the test medium is observed.

II Reagents and lab wares Inoculating wire, Bunsen flame, test tubes, fungi from culture tube, and Christensen’s urea agar slope.

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KEY FACTS 1 Certain fungi possess the enzyme urease that hydrolyzes urea releasing ammonia into the medium. 2 Phenol red that is incorporated in the medium changes its color from yellow to red in alkaline pH, thus indicating the presence of urease activity. 3 Always check the sterility of the slants before inoculation. 4 An un inoculated medium must be incubated along with the test. 5 Observe the growth of inoculum irrespective of change in colour.

VIVA 1 What is the medium used in urease test? 2 What is the principle of urease test? 3 Give examples of urease positive fungi?

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000.

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LESSON

79

Carbohydrate Assimilation Test

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to:

1 Pipette 2.0 ml. of sterile saline into a test tube. 2 With a sterile loop pick up few isolated colonies of the organism from the SDA plate and emulsify to a turbidity equal to McFarland 4 units. 3 Cover the surface of yeast nitrogen base agar with the suspension of the yeast cells. 4 Remove the excess fluid and allow the surface of the agar to dry. 5 With sterile forceps place the carbohydrate disc onto the surface of agar in such a way that at least 30 mm space is present between each disc. 6 Incubate the plate at 30°C or 37°C for 24–48 hours. 7 At the end of the incubation period observe the plate for growth around the disc.

1 Find out the pattern of assimilation of carbohydrates by yeast and yeast-like fungi.

INTRODUCTION Yeast and yeast-like fungi use carbohydrates as sources of energy. Hence in this test, utilization of carbohydrate is used as a definitive diagnosis for yeast or yeast-like fungi.

PRINCIPLE Yeast and yeast-like fungi are identified by the pattern of carbohydrate assimilation. They are inoculated on the carbohydrate-free yeast nitrogen base agar on which different filter paper discs containing various carbohydrates are placed. After incubation for appropriate time, growth around the discs is observed and the carbohydrate utilization pattern is assessed.

QUALITY CONTROL 1 Carbohydrate discs and media should be checked using standard control strains as follows: Candida albicans ATCC 14053 C. guilliermondii ATCC 6260 C. pseudotropicalis ATCC 4135 2 Sterility of glassware and media should be ensured.

REQUIREMENTS

OBSERVATION

I Equipments Incubator.

After incubation, growth of the fungi around the discs is observed and the carbohydrate utilization pattern is assessed.

II Reagents and lab wares Standard laboratory glassware and equipments, filter paper discs, McFarland standard, yeast nitrogen base agar (Table 79-1), sterile saline, and filter paper discs containing different carbohydrates III Specimens Pure growth of test fungi on Sabouraud’s dextrose agar (SDA) medium.

RESULTS AND INTERPRETATION Growth of the fungi around each carbohydrate disc is observed and results are interpreted in such a way that if the fungus has grown touching the edges of the disc, and that organism assimilates that particular carbohydrate in the disc then it is considered positive. Similarly, if the fungus does not grow in the proximity of a particular carbohydrate disc, it is considered negative for assimilation of that carbohydrate.

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Table 79-1 Preparation of Yeast Nitrogen Base Composition

Preparation

Boric acid – 500 µgm Copper sulphate – 40 µgm Potassium iodide – 100 µgm Ferric chloride – 200 µgm Manganese sulphate – 400 µgm Sodium molybdate – 200 µgm Zinc sulphate – 400 µgm Biotin – 2 µg Calcium pantothenate – 400 µgm Folic acid – 2 µg Inositol – 2000 µgm Niacin – 400 µgm p-aminobenzoic acid – 200 µgm Pyridoxine hydrochloride – 400 µgm Riboflavin – 200 µg Thiamine hydrochloride – 400 µgm L-Histidine monohydrochloride – 10.0 milligram DL-Methionine – 20 mg DL-Tryptophan – 20 mg Magnesium sulphate – 500 mg Sodium chloride – 100 mg Ammonium chloride – 5 g Monopotassium phosphate – 1 g Distilled water – 1000 ml

All the above ingradients are to be sterilized by filtration and dispersed aseptically. Add. 10 ml. of the above solution to 90ml of 2% molten agar and pour into the plates.

KEY FACTS 1 Turbidity using the organism should be done in such a way that it is equal to McFarland Standard 4 units. 2 Sterility should be maintained for medium and carbohydrate discs. 3 Growth of fungi near carbohydrate discs should be confirmed after thorough examination.

VIVA 1 Describe how carbohydrate medium is prepared. Ans. The basic salt solution is prepared and 10 ml. of this solution is added to 90 ml of 2% molten agar and poured in plates. 2 How do you interpret carbohydrate assimilation test result?

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000. pp. 232.

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LESSON

80

Carbohydrate Fermentation Test

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Demonstrate the ability of different yeast and yeast-like fungi to ferment various carbohydrates.

INTRODUCTION Yeasts and yeast-like fungi use carbohydrates as sources of energy. Fermentation of carbohydrate in the medium produces a color change and bubbles which can be used as a marker to find out the type of yeast in culture.

Preparation of indicator broth medium: The indicator broth medium (IBM) contains peptone, 1.0 gm.; sodium chloride, 0.5 gm.; beef extract, 0.5 gm; bromocresol purple (0.4%),10 ml and distilled water, 90.0 ml. Durham’s tube is placed in an inverted position in screw capped test tubes. In each tube, fill 5–6 ml IBM just above the Durham’s tube and sterilize at 121°C for 15 min. Aseptically, 0.3 ml of 20% filter sterilized carbohydrate solution (glucose, lactose, sucrose, maltose, etc.) are added to the medium. III Specimen Pure growth of test fungus on Sabouraud’s dextrose agar (SDA) medium.

PROCEDURE PRINCIPLE Sometimes, a definite identification of the suspected fungi cannot be made using carbohydrate assimilation test. In such cases, carbohydrate fermentation tests are used as a supplement. Fermentation of carbohydrates produces a visible color change in the medium, and bubbles will be produced in the Durham’s tube. This is an indicator of fermentation. Glucose, maltose, sucrose, lactose, galactose, and trehalose are the sugars used in carbohydrate fermentation tests.

1 Take tubes with indicator broth medium with sugars. 2 With a sterile loop pick up few isolated colonies of the fungus from the SDA plate and emulsify to a turbidity equal to McFarland 4 units. 3 Inoculate each sugar tube with 0.2 ml (5 drops) of culture suspension. 4 Incubate the tubes at 30°C or 37°C for 2-10 days. Note: Do not screw caps on tubes tightly. 5 Observe the presence of air bubbles in Durham’s tubes.

REQUIREMENTS

QUALITY CONTROL

I Equipments Incubator. II Reagents Standard laboratory glassware and equipments, McFarland standard. Different carbohydrates, and indicator broth medium (IBM).

1 Quality of sterile glassware must be checked for proper sterility. 2 Sterility of medium used should be confirmed before use. 3 Quality of medium used should be checked by growing known standard strains of fungi. Candida albicans: Glucose, maltose positive. Candida kefyr: Glucose, sucrose positive.

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OBSERVATIONS

RESULTS AND INTERPRETATION

After incubation, the Durham’s tubes are observed for presence of any gas bubbles.

Presence of bubbles or drop in the fluid level in Durham’s tube indicates fermentation of sugars. Development of yellow color is not a reliable indicator of fermentation and is ignored.

KEY FACTS 1 Carbohydrate fermentation test is a supplementary test only when there is difficulty in making a definitive identification using carbohydrate assimilation test. 2 Development of yellow color is not reliable indicator of fermentation and hence production of air bubble in Durham’s tube is to be noted.

VIVA 1 2 3 4

What are the different carbohydrates used in carbohydrate fermentation test? What percentage of carbohydrate should be incorporated in carbohydrate medium? How do you interpret carbohydrate fermentation test? What are the advantages of carbohydrate fermentation test over carbohydrate assimilation test?

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology. Interprint. 2000.

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LESSON

81

Identification of Common Fungi

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to:

Lactophenol cotton blue wet mount preparation of the fungus (refer chapter 73).

1 Know important characteristic morphological features of different fungi. 2 Identify a given fungus.

QUALITY CONTROL Quality of sterility of medium should be checked before use.

INTRODUCTION OBSERVATION

Fungi cause varied infections in man, hence identification of the fungi is a very important approach in patient management. Hence it is necessary to know different fungi causing infections in man and methods to identify them.

The wet mount preparation should be visualized under low power (10x) and high power (40x) objectives and the morphology observed.

PRINCIPLE

RESULTS AND INTERPRETATION

In a mycology laboratory, the fungi can be cultivated, and its colonial characteristics will throw some light on the type of fungus. Further, the fungal element can be stained and observed under a microscope and presumptive identification can be done.

REQUIREMENTS I Equipments BOD incubator and dissecting microscope.

II Reagents and lab wares Standard laboratory glassware, Bunsen burner, hard lens, glass slides, cover slips, sterile cotton swab, glass marking pencil, SDA medium, lactophenol cotton blue stain. III Specimen Fungal culture grown on SDA’s medium.

By microscopic observation, characters of the yeast, mold, hyphae, conidia, etc. can be made out and used in classifying the type of fungus isolated. Molds Rhizopus Mucor Alternaria Fusarium Aspergillus Penicillium Cladosporium Cephalosporium Trichophyton Epidermophyton Yeast Torulopsis Candida

Rhizopus A Common laboratory contaminant. Colony morphology Rapidly growing white colored fungus swarms over entire plate showing aerial mycelium cottony.

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Identification of Common Fungi

Microscopy Spores are oval, colorless and sometimes brown. Mycelium non-septate, sporangiophores straight and terminate in black sporangium containing sporangiospores and columella; root like hyphae (rhizoids) penetrating the medium.

Mucor Common food contaminant. Colony morphology Resembles colony of Rhizopus. Microscopy Oval spores, non-septate mycelium giving rise to single sporangiophores and terminate in globular sporangium containing columella and sporangiospores.

Alternaria Usually found on plants. Colony morphology Grayish green or black colonies with grey edges and swarming over entire plate with grayish white aerial hyphae. Microscopy Conidia multi celled, pear-shaped and attached to single conidiophores arising from septate mycelium.

Fusarium Usually found in soil. Colony morphology Woolly white colonies may change to pink, purple or yellow (Fig. 81-1). Microscopy Conidia multi celled, oval or crescent shaped, attached to conidiophores arising from a septate hypha.

Aspergillus Usually infect plants and animals. More than 170 species are known. Almost sixteen different species cause human disease. They are: Aspergillus fumigatus Aspergillus flavus Aspergillus niger Aspergillus oryzae Aspergillus amstelodami Aspergillus verscicolor Aspergillus terreus Aspergillus nidulans Aspergillus candidus Aspergillus ustus Aspergillus carnens Aspergillus arenaceus Aspergillus clavatus Aspergillus caseillus Aspergillus restricutus The characteristics used to identify them are: A) Colour and shape of conidial head. B) Number of sterigmata/phialids. C) Shape of vesicles. D) Colour of conidiophore. E) Presence and absence of cleistothecia, and F) Size and shape, and color of ascospore and conidia.

Aspergillus fumigatus Colonies Colonies are blue-green, velvety, surface powdery or granular, sometimes radially folded (Fig. 81-2). Conidial head Columnar, compact. Conidiophore Smooth, terminate into club or flask shaped green. Vesicle Club or flask shaped, green. Sterigmata Produced on upper half of the vesicle, uniseriate, appears green in color and crowded. Axis of sterigmata is roughly parallel to that of conidiophore and conidia produced in chains on sterigmata.

FIGURE 81-1 SDA with colonies of Fusarium species.

Conidia Globose, green and echinulate.

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FIGURE 81-2 SDA with colonies of Aspergillus fumigatus.

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FIGURE 81-4 SDA with colonies of Aspergillus flavus.

Aspergillus niger Aspergillus flavus Colonies White, fluffy, reverse buff colored, covered with black spores (Fig. 81-3). Conidial head large black to brownish, initially globose, become radiate then splits into divergent spore columns. Conidiophore Variable in size, thick smooth walls, conidiophore hyaline and brownish near the vesicle. Vesicle Globose, concave under surface, brownish sterigmata produced in two series, septate primary sterigmata and short secondary sterigmata. Conidia Globose, echinulate.

Colonies Rapidly growing, yellowish green and velvety colony (Fig. 81-4). Conidial head Radiate, loosely columnar. Conidiophore hyaline, thick walled and roughened. Vesicle Globose, sub globose or elliptical and sterigmata cover entire surface or at least three fourth. Sterigmata Monoseriate and biseriate sterigmata seen in same strain, produced by small and large vesicles respectively. Conidia Conidia globose, subglobose or elliptical. Conidia echinulate and appears yellow green.

Penicillium species Colonies Initially white and fluffy, later turns to green and blue green, with radial folds. Hyphae Septate, hyaline. Conidiophores Long with branching phialids.

FIGURE 81-3 SDA with colonies of Aspergillus niger.

Phialids Flask shaped, branched and gives rise to brush like appearance, producing sterigmata.

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Identification of Common Fungi

Sterigmata Long with tapering end, producing chains of conidia. Conidia Long chains of conidia, spherical or oval in shape.

Macroconidia String-like, bean shaped, septate, with characteristic rat tail appearance, rarely produced.

Cladosporium

Microconidia Tear-shaped, rarely produced. Swollen antler-like hyphae produced rarely, may produce chains of chlamydospores.

Usually found on dead and decaying plants.

Trichophyton violaceum

Colony Small, heaped, greenish black and powdery.

Colony Slow growth, port wine or deep violet colored, flat or heaped up, surface waxy and glabrous, wrinkled loose pigment present.

Microscopy Conidia develop at the end of complex conidiophores arising from brown septate mycelium.

Macroconidia Generally not present. Thick walled structures resembling macroconidia of T. rubrum may be present.

Cephalosporium Used in antibiotic production Colony Rapidly growing compact, moist colonies, cottony, with gray or rose colored aerial hyphae. Microscopy Single celled conical or elliptical conidia held together in clusters at the tips of conidiophores. Erect, unbranched conidiophores arise from septate mycelium.

Trichophyton verrucosum Colony Slow grower, white glabrous heaped up colony, sometimes button like with velvety texture. No pigment produced on reverse. Variants with yellow or grey white colonies, rugal folds seen.

Microconidia Generally not present, pyriform. Chains of chlamydospores produced.

Epidermophyton floccosum Colony Velvety or powdery surface, surface folded in radiating furrows, reverse of colony yellow tan. Macroconidia Clavate, smooth, thick walled, septa 2–3, characteristics clusters of twos and threes. Microconidia Not produced. Chlamydospores Produced, racquet hyphae and nodular bodies present.

KEY FACTS 1 Characters of aerial hyphae, septation, conidial shape, size and color are very important for identifying the fungus. 2 Colony characters, color, obverse and reverse should be kept in mind.

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VIVA 1 What are the common fungi that cause disease in humans? Ans. The common fungi that cause human infections include pathogenic species of Curvularia Rhizopus Mucor Alternaria Fusarium Aspergillus Penicillium Cladosporium Cephalosporium Trichophyton Epidermophyton Torulopsis Candida Absidia Cryptococcus Histoplasma Sporothrix Blastomyces, etc 2 How are culture and staining of fungal elements useful in characterizing fungi? Ans. Different fungi have their own predilection to grow on different media depending upon the exact nutritional requirement. Moreover different fungi produce a wide variety of morphological structures like conidiophores, conidia, sporangiophores, sporangia, septate or aseptate hyphae, etc., which on staining can be visualized and the fungi identified. 3 What are different types of spores or conidia produced by different molds? Ans. Different molds produce different types of spores and conidia, like chlamydospores, ascospores, microconidia, macroconidia etc. 4 How is the colony morphology of fungi important in diagnosing fungal infections? Ans. The colonies produced by many pathogenic fungi differ greatly from each other. Moreover production of pigments by these colonies also vary greatly. Hence this helps for a preliminary identification of the type of fungus

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s Microbiology and Microbial Infections. 9th Edition. Mycology. Volume 4. pp. 711. Arnold publishers. 2 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 3 Jagadish Chander. A Text Book of Medical Mycology.. Interprint. 2000.

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UNIT

XI Virology

Lesson 82

Cultivation of Viruses in the Cell lines

Lesson 83 Cultivation of Viruses in Embryonated Egg

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LESSON

82

Cultivation of Viruses in the Cell lines

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know and understand the common cell lines used for routine maintenance of viruses in a virology laboratory.

INTRODUCTION Viruses are intracellular pathogens which are dependent on host cell machinery for their multiplication and growth. Now a days monolayer cell cultures are mostly used in diagnostic and research work in viral diseases. Cell culture is the most widely used system for cultivation of viruses since it is more convenient method compared to the other methods like egg inoculation and animal inoculation. The cell cultures are used for a) isolation of viruses from clinical specimens for diagnosis of viral diseases, b) biochemical studies of viral replication and c) production of viral antigens and vaccines. Depending upon the number of divisions which a cell line undergoes in-vitro before dying, the cell lines have been classified as primary, diploid and continuous cell lines. The most commonly used cell culture systems in most laboratories include chick embryo fibroblast cells, human amnion cells, Rhesus monkey kidney cells, HeLa (Human carcinoma of cervix cell line), Hep 2 (Human epithelioma of larynx cell line), McCoy (Human synovial carcinoma cell line),Vero (Vervet monkey kidney cell line), and W1-38 (Human embryonic lung cell line).

Many viruses kill the infected viral cells in which they grow and bring about detectable changes in morphology of the cells. These changes are collectively known as cytopathic effects. Some viruses however do not produce any cytopathic effect (e.g., rubella virus). The most important precaution to be taken during maintenance of cell lines is sterility. Contamination of cell lines should be prevented and even cross contamination among cell lines should be avoided.

REQUIREMENTS I Equipments Inverted microscope (Fig. 82-1), incubator (Fig. 82-2), haemocytometer and biological safety cabinet. II Reagents and lab wares Sterile glassware, pre-sterilized tissue culture plasticware (Fig. 82-3), Pasteur pipettes and measuring pipettes, membrane filter, syringes, vials, discard jar, Eagle’s, minimum essential medium (MEM), sodium bicarbonate (NaHCO3), EDTA trypsin mixture, foetal calf serum (FCS), sterile double distilled water, virus inoculum, spirit and sodium hypochlorite. Monolayer of a cell culture in a culture flask is treated with trypsin or versene to disperse cells. III Specimen Suspected virus infected specimen like the cerebrospinal fluid (CSF), stool, rectal swab, and throat swab.

PRINCIPLE

PROCEDURE

Viruses infect healthy cells grown in the laboratory. When susceptible cells are used for inoculation of viruses, they show pathological changes and the viruses can be harvested from the cells for further tests. The growth of viruses in the cell lines can be known by a) cytopathic effects, b) immunofluorescence, c) haemagglutination and haemadsorption, and d) interference.

1 Discard the trypsin versene mixture and add a small amount of MEM with 10% FCS to the monolayer of cells. 2 Count the cells with the medium in a hemocytometer for appropriate splitting. 3 Inoculate the cells into sterile flasks or tubes for viral inoculation.

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4 Fill the new flasks with MEM and incubate in horizontal position. 5 Select a healthy monolayer, which is also confluent, for viral inoculation. 6 Inoculate the monolayer of cells with virus using sterile Pasture pipette, and incubate at 37°C. 7 Observe for the cytopathic effect (CPE) 7 days after inoculation.

QUALITY CONTROL

FIGURE 82-1 Inverted microscope.

1 Sterility precautions should be taken perfectly. 2 Susceptible cells to be selected for the appropriate virus.

OBSERVATIONS After incubation, the flasks are observed for confluency and healthy monolayer of cells and virus infected cells are classified. Viruses are known to produce cytopathic effects are identified by observing the same in the infected cell lines. Non-cytopathogenic viruses are identified by other methods like immunofluorescence, haemagglutination and haemadsorption, and interference

FIGURE 82-1 Incubator.

RESULTS AND INTERPRETATION The cell lines are observed for any cytological alterations that are diagnostic of viral infections (Table 82-1).

FIGURE 82-3 Cell culture bottle.

Table 82-1 The cell lines and indications for the viruses and cytopathic effects they produce Type of viruses

Susceptible cell line

Cytopathic effect

Adenovirus

Primary human embryonic kidney cells. HeLa cells. HEp 2 cells. Human kidney cells. Human fibroblast cells. Hep 2 cells. Vero cells. RD cells. Primary human and monkey kidney cells. LLC-MK2.

Rounding and clustering of swollen cells.

Herpes simplex virus Varicella zoster virus Cytomegalovirus Enterovirus

Paramyxovirus

Rounding and swelling of cells. Multinucleated giant cell formation. Giant cell formation. Rapid rounding of cells progressing to complete cell destruction. Syncytia formation.

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KEY FACTS 1 The most important aspect to be taken care of in cell culture is sterility. Hence precautions for sterility should be meticulously followed. 2 Susceptible cells should be selected for the appropriate viral inoculation.

VIVA 1 Why is cell culture method, the most preferred method in a virology laboratory? 2 What precaution is to be taken during maintenances of cell lines? 3 What are the commonly used cell lines in virology laboratory?

FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s; Microbiology and Microbial Infections. 9th Edition. Virology. Volume 1. Arnold publishers. 2 Knipe DM, Howley PM. Field’s Virology. 4th Edition. Lippincott Williams and Wilkins. 2001. 3 Zuckerman AJ, Banatvala JE, Pattison JR, Griffiths PD and Schoub BD. Principles and Practice of Clinical Virology. 5th Edition. John Willey and sons Ltd. 2004.

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LESSON

83

Cultivation of Viruses in Embryonated Egg

LEARNING OBJECTIVES

1 Know the routes of inoculation of eggs for propagation of viruses.

Allantoic cavity is mainly employed for harvesting influenza virus for vaccine production , and also for yellow fever (17D strain) and rabies (Flury strain) vaccines. CAM is mainly used for growing pox viruses. These produce characteristic visible lesions such as pocks. Pocks produced by different viruses vary in their morphology.

INTRODUCTION

REQUIREMENTS

Prior to 1950s the technique of propagation of viruses in embryonated eggs was popular because of non-availability of cell culture techniques during those times. The eggs are used because they are inexpensive, easily available and much simpler to handle than animals. The viruses can grow in different compartments of the egg . Since eggs lack a well developed defence mechanism of their own, they do not interfere with growth of viruses. However, many viruses fail to grow on primary inoculation into eggs. Nevertheless, the embryonated eggs are still used for the isolation of avian viruses, influenza viruses and also for vaccine production.

I Equipments Egg holders, candling lamp and hole puncher,

After completing this practical you will be able to:

PRINCIPLE Usually 8-11 days old chick embryos are used. Age of the egg chosen depends on route of inoculation described since various membranes and their contents vary in size as embryo matures. These eggs are inoculated by one of the following routes: yolk sac, amniotic sac, allantoic cavity and chorioallantoic membrane (CAM). Any of the viruses or specimen suspected to contain viruses are inoculated. After inoculation, eggs are incubated for 2-9 days. Viral growth is recognized by death of embryo, formation of pocks, or haemagglutination property of embryonic fluid. Yolk sac is mainly used for culture of some viruses, certain bacteria (Chlamydia and Rickettsia) and parasite (Toxoplasma gondii). Amniotic sac is used for the primary isolation of influenza virus.

II Reagents and lab wares Syringes (1 tuberculin syringe preloaded with 0.1 ml of 10-3 dilution of NDV in GLB), gauze, pencils, gloves, sterile forceps, Pasteur pipette with bulb, screw capped sterile vials, egg, melted paraffin wax and 70% ethanol. The eggs are candled to determine the position of the embryo and its viability. Since viruses need living tissues to replicate, candling is important. When candled, healthy embryo has an orange color with evident vasculature. Dead embryo shows clear yellow with no vasculature. Black, green or brown color indicates contamination. Dead embryos are promptly discarded. III Specimen Virus isolate or specimen suspected to contain virus.

PROCEDURE 1 Disinfect the egg shell. 2 Drill the egg shell. 3 Inoculate the specimen in the embryonated egg through appropriate route. Note: Age of the embryonated egg is chosen depending on the route of inoculation since various membranes and their contents vary in size as embryo matures (Table 83-1). 4 After 2–5 days post injection, viral growth in the egg is recognized by death of embryo, pocks, or hemagglutination.

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QUALITY CONTROL

RESULTS AND INTERPRETATION

1 Before inoculation, the eggs are candled to determine the position of the embryo and its viability. 2 Age of the embryonated egg is chosen depending on the route of inoculation since various membranes and their contents vary in size as embryo matures. 3 Sterile precautions should be taken throughout the procedure.

Viral growth in the inoculated egg is recognized by death of the embryo, pock formation in CAM (Fig. 83-1), haemagglutination, etc.

OBSERVATIONS The inoculated part of embryonated egg is observed for changes due to viral infection. FIGURE 83-1 CAM showing pocks.

Table 83-1 The routes of inoculation of the egg and the viruses isolated Route of inoculation

Age of embryo

Virus

Use

Chorio allantoic membrane.

10-14 days.

Variola virus. Herpes simplex virus. Vaccinia virus.

Isolation. Isolation and typing. Vaccine titration.

Amniotic sac.

10-12 days.

Influenza A virus. Mumps virus.

Isolation. Isolation.

Allantoic sac.

9-12 days.

Mumps virus. Influenza A, Influenza B. Parainfluenza viruses.

HA antigen preparation. Hybrid vaccine production. Isolation.

Yolk sac.

6-8 days.

Flavi viruses.

Isolation, vaccine production.

KEY FACTS 1 Usually 8-11 days old chick embryos are used for cultivation of viruses. 2 Since viruses need living tissues to replicate, candling of eggs is important.

VIVA 1 Why is the age of the embryo an important factor in virus inoculation? 2 What are some important parameters to be noted before passaging viruses in embryonated eggs? 3 What is the purpose of candling? Ans. Since viruses need living tissues to replicate, candling is important. When candled, healthy embryo has an orange color with evident vasculature, dead embryo show clear yellow colour with no vasculature, black, green or brown color indicates contamination. 4 Would you expect a higher yield of virus from dying or dead embryo, why? Ans. Higher yield of viruses cannot be expected from dying or dead embryo since viruses require healthy living tissues for their multiplication.

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FURTHER READINGS 1 Collier L, Balows A, Sussman M. Topley and Wilson’s; Microbiology and Microbial Infections. 9th Edition. Virology. Volume 1. Arnold publishers. 2 Knipe DM, Howley PM. Field’s Virology. 4th Edition. Lippincott Williams and Wilkins. 2001. 3 Zuckerman AJ, Banatvala JE, Pattison JR, Griffiths PD and Schoub BD. Principles and Practice of Clinical Virology. 5th Edition. John Willey and sons Ltd. 2004.

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UNIT

XII Microbiology of Water, Milk and Air

Lesson 84 Microbiology of Water Lesson 85 Microbiology of Milk Lesson 86 Microbiology of Air

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LESSON

84

Microbiology of Water

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know various methods used for testing potable water. 2 Test the quality of the potable water for drinking.

The total coliform count is widely regarded as the most reliable indicator of potable water quality. However, the presence of coliforms not necessarily indicate faecal or sewage contamination, because, these organisms are present in large quantities in the environment.

Faecal or thermotolerant coliforms INTRODUCTION Drinking water is acceptable and fit for drinking when it is clear, colourless, odourless and without disagreeable taste. Microscopically it should be free from pathogenic organisms. Natural sources of water usually contain some saprophytic bacteria, such as Pseudomonas, Serratia, Flavobacterium, Chromobacterium, Acinetobacter, Alcaligenes, etc. These saprophytes are harmless. Water gets contaminated by pathogens which are introduced into water by sewage pollution. A wide varieties of diseases are transmitted by contaminated water. There are some indicator organisms, whose presence indicates the contamination of water with fecal matter. These are: 1 2 3 4 5

Coliforms. Fecal (thermotolerant) coliforms. Faecal Escherichia coli. Faecal Streptococci, and Clostridium perfringens.

Coliforms Coliforms are defined as members of the family Enterobacteriaeceae which grow in the presence of bile salts and produce acid and gas from lactose within 48 hours at 37°C. In order to include anaerogenic bacteria and those which are nonlactose fermenters, it has been modified as the members of the Enterobacteriaeceae capable of growing galactosidase at 37°C that normally posses.

These satisfy the criteria for coliforms but have the additional property of the ability to grow at a higher temperature 44°C.

Faecal Escherichia coli These are defined as thermotolerant coliforms which ferment lactose (or) mannitol at 44°C with the production of acid and gas within 24 hours and also form indole from tryptophan at 44°C. The presence of E. coli is considered as the contamination of water with feces of human or animal origin.

Faecal streptococci These are catalase negative, Gram positive cocci present in the intestinal tract of man and animals. These organisms have the Lancefield group D antigen, hydrolyse aesculin and can grow at 45°C, in the presence of azide and 40% bile. Such organisms which can survive 60°C for 30 min and can grow at 10°C at pH 9.6 and in 6.5% of sodium chloride (NaCl) belong to the genus Enterococcus (Enterococcus faecalis and E. faecium) .

Clostridium perfringens The presence of this organism in water in the absence of other indicators of contamination of water implies remote or intermittent fecal pollution. Viruses in water are destroyed by chlorination. When the concentration of free residual chlorine is at least 0.5 mg per litre, for a minimum contact period of 30 minutes at pH below 8 and a turbidity of 1 nephelometric turbidity unit or less, protozoa such as Entamoeba histolytica, Giardia species and

Textbook of Practical Microbiology

Balantidium coli may be present in the drinking water. Coliforms are not the reliable indicator of protozoal contamination.

Collection of water samples Water is collected in heat sterilised bottles containing a sufficient volume of sodium thiosulphate to neutralise the bactericidal effect of any chlorine or chloramine in the water. For collection from tap, water is allowed to run to waste for 2–3 min. before collecting it into the bottle. When collecting water samples from lakes or streams the bottle is opened at a depth of about 30 cm with its mouth facing the current. At least 100 ml of water to be tested is collected in each bottle. After collecting the water in the bottle, the bottle is stoppered, and sent to the laboratory as quickly as possible within 6 hours keeping it in a cool container and protecting it from light.

PRINCIPLE The following tests can be done for bacteriological analysis of water: 1 Plate count. 2 Detection of coliform bacteria and E. coli. a Presumptive coliform count: multiple tube technique. b Differential coliform test. c Membrane filtration method. 3 Detection of faecal streptococci. 4 Examination of Cl. perfringens, and 5 Test for pathogenic bacteria.

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Differential coliform test This is called Eijkman test. This test is usually employed to find out whether the coliform bacilli detected in the presumptive test are E. coli or not. After the usual presumptive test, subcultures are made from all the bottles showing acid and gas to fresh tubes of single strength MacConkey medium already warmed to 37°C. They are incubated at 44°C for 24 hours. Those showing gas in Durham’s tubes contain E. coli. From the number of positive tubes obtained, results are read off the probability tables. Further confirmation of the presence of E. coli is done by testing for indole production and citrate utilization tests.

Membrane filtration method A measured volume of water is filtered through a Millipore filter. All the bacteria present are retained on its surface. This is then placed on suitable media and incubated at the appropriate temperatures. Colonies on the surface of the membrane are counted. After 18 hours of incubation the presumptive coliform counts and detection of E. coli can be directly made.

Detection of faecal streptococci Subcultures are made into tubes containing 5 ml of glucose azide broth from the positive bottles in the above test. Streptococcus faecalis if present produces acid in the medium within 18 hours at 45°C.

Examination for Cl. perfringens Water is incubated in litmus milk medium at 37°C for 5 days, if positive, stormy fermentation occurs.

Examination for pathognic bactrial cells test This consists of inoculating the nutrient agar with water to be tested and incubating the agar aerobically in parallel at 37°C for 1–2 days and at 22°C for 3 days. After incubation number of the colonies formed in the agar are counted. Those which grow at 37°C are associated with organic material of human or animal origin and those growing at a lower temperature are mainly saprophytes that normally inhabit water, soil, and vegetables. The agar count at 22°C gives an indication of the amount of decomposing organic matter in the water available for bacterial nutrition. Though these are non-pathogenic the greater the amount of organic matter present, the more likely is the water to be contaminated with parasitic and potentially pathogenic organisms. The agar count at 37°C is a more important index of dangerous pollution.

Detection of coliform bacteria and E.coli Presumptive coliform test – Multiple tube technique The test is called presumptive, because of the presumption that the reactions are due to coliforms organisms. The count is made by adding varying quantities of water (0.1 ml–50 ml) to bile salt lactose peptone water with an indicator for acidity. Acid and gas formation indicate the growth of coliform bacilli.

For other special pathogens like Salmonella, Vibrio, etc. corresponding special media are used. In this chapter Presumptive coliform test and diffrential coliform test will be discussed.

REQUIREMENTS I Equipments Bacteriological incubator, autoclave and water bath. II Reagents MacConkey broth, brilliant green bile broth Preparation of MacConkey broth: This is prepared by mixing pancreatic digest of gelatin, 20 grams; lactose, 10 grams; bile salt, 5 grams; bromocresol purple, 0.01 grams and final pH (at 25°C) is adjusted at 7.3 ± 0.2. Preparation of double strength medium: It is prepared by suspending 35 grams in 500 ml of distilled water and heated to dissolve the medium completely. 50 ml and 10 ml of the media are dispensed into screw capped bottles with inverted Durham’s tubes and are sterilized by autoclaving at 15 lbs (121°C for 15 minutes). Preparation of single strength medium: It is prepared by suspending 3.5grams in 100 ml of distilled water. 5 ml quantities of the medium are dispensed in screw capped bottles with

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inverted Durham’s tubes and are sterilized by autoclaving at 15 lbs (121°C for 15 minutes). Preparation of brilliant green bile broth (BGBB): This is prepared by mixing peptic digest of animal tissue, 10 grams; lactose, 10 grams; ox gall, 20 grams and brilliant green, 0.0133 gm and adjusting the final pH (at 25°C) to 7.2 ± 0.2. 4 grams of the medium is suspended in 100 ml distilled water and mixed well. The media is distributed in 5ml volumes in test tubes with inverted Durham’s tubes and sterilized by autoclaving at 15 1bs pressure (121°C) for 15 minutes. III Specimen Water specimen to be tested.

PROCEDURE 1 Collect 500 ml of water in a sterile bottle. 2 Inoculate water sample immediately into the MacConkey broth medium. 3 Inoculate 50 ml of water into 50 ml double strength media (1 bottle) 4 Inoculate 10 ml of water into 10 ml double strength medium (5 bottles). 5 Inoculate 1 ml of water into 5ml single strength medium (5 bottles). 6 Incubate all the bottles at 37°C for 48 hours. 7 Check for acid and gas after 24 hours and 48 hours. 8 Any positives are inoculated into brilliant green bile broth and peptone water. 9 Incubate at 44°C in a water bath overnight.

QUALITY CONTROL Sterility of media and strength of media should be properly checked.

OBSERVATIONS Observe for the presence of gas and production of indole. Presence of gas in brilliant green bile broth and indole production at 44°C is indicative of the presence of E. coli. For interpretation refer MacGrady’s table.

RESULTS AND INTERPRETATION Presumptive coliform count The no. of bottles showing acid and gas is counted and compared with the MacGrady’s table. Result is expressed as- coliforms most probable no (MPN)/ 100ml. e.g. for Presumptive coliform count No. of tubes giving positive reaction 50 ml 10 ml 1 ml 1 2 2 Result: Coliforms most probable no 10/100 ml. Differential coliform count The no of tubes showing positive results i.e. both gas productions in BGBB and Indole test at 44°C is compared with MacGrady’s table. Result is expressed a Escherichia coli Most probable No (MPN) / 100 ml. Eg. for Differential coliform counts: No. of tubes giving positive reaction. 50 ml 10 ml 1 ml 1 1 1 Results: Escherichia coli MPN —— 5 /100 ml Report: Unsatisfactory. Grades of the quality of drinking water supplied as determined by results of periodic Escherichia coli and coliform counts is listed in the table 84-1.

Table 84-1 Grades of the Quality of Drinking Water Supplies Determined by results of Periodic Escherichia coli and Coliform counts Quality of supply

Results from routine samples Coliform counts / 100 ml E. coli count / 100 ml

Tolerance

1 Excellent. 2 Satisfactory.

0 1–3

In all samples. Provided that coliforms do not occur in Consecutive samples or in more than 5% of samples.

3 Intermediate. 4 Unsatisfactory.

4–9 0 10 coliforms or 1 or more E. coli or any coliform present in consecutive samples, or presence of any coliform organisms in more than 5% of routine samples.

0 0

In any sample.

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KEY FACTS 1 Drinking water is acceptable and fit for drinking when it is clear, colourless, and odourless and without disagreeable taste. Microscopically it should be free from pathogenic organisms. 2 Coliforms are defined as the members of the family Enterobacteriaeceae which grow in the presence of bile salts and produce acid and gas from lactose within 48 hours at 37°C. 3 They are indicator of faecal contamination. 4 Coliforms are not the reliable indicator of protozoal contamination. 5 Eijkman test is usually employed to find out whether the coliform bacilli detected in the presumptive test are E. coli or not.

VIVA 1 What are the common water-borne infections transmitted by contaminated water? Ans. Typhoid fever, cholera, poliomyelitis, viral hepatitis, amoebiasis, giardiasis, cysticercosis, etc. 2 Mention the routine laboratory tests done for the analysis of water. 3 Which are the organisms which indicate fecal contamination of water?

FURTHER READINGS 1 Ananthanarayanan. R, Paniker. C.K, Ananthanarayanan and Paniker’s Textbook Of Microbiology; 7th Edition, Orient Longman, pp 603 – 609, 2005. 2 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996.

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LESSON

85

Microbiology of Milk

LEARNING OBJECTIVES After completing this practical you will be able to:

number of viable bacteria in the milk. For un refrigerated milk there is a consistent relationship between the bacterial count and the dye reduction time.

1 Test the quality of the un refrigerated milk by methylene blue test.

REQUIREMENTS

INTRODUCTION

I Equipments Water bath.

Human infections may be caused by the ingestion of animal milk which contains microorganisms derived either from the animal (faeces), from the environment, or from milk handlers such as dairy workers. Certain precautions are taken to prevent contamination. A wide number of bacteria are found in contaminated milk (Table 85-1). The infections that can be transmitted from infected animals to humans through contaminated milk are many. They include tuberculosis, brucellosis, streptococcal and staphylococcal infections, salmonellosis and Q fever. The routine bacteriological examination of milk to detect bacterial contaminations can be done by the following methods: 1 2 3 4 5 6

Viable count. Test for coliform bacilli. Methylene blue reduction test. Phosphatase test. Turbidity test, and Examination for specific pathogens.

Of these tests, methylene blue test is a rapid, simple, inexpensive and most widely used test for testing the milk. This method will be described in this chapter.

PRINCIPLE Time taken for bacterial dehydrogenases to reduce methylene blue dye and decolourise it, is taken as an indicator of the

II Reagents and glass wares Sterile test tubes, sterile pipettes, sterile rubber stopper, methylene blue tablets and distilled water. III Specimen Milk to be tested.

PROCEDURE The time of reduction is affected by the temperature at which the milk is held before testing. The test should not be done if the atmospheric shade temperature exceeds 21°C. 1 Prepare a 1 in 3,00,000 solution of methylene blue by dissolving a standard methylene blue tablet in 200ml cold sterile glass-distilled water and making upto 800ml with more of such water. Note: This solution can be stored in dark in a refrigerator and can be used for 2months. 2 Mix thoroughly the milk sample. 3 Aseptically transfer 10ml of sample of milk with a pipette to a sterile test tube. 4 Add 1ml of methylene blue solution by a 1ml sterile pipette. 5 Put a sterile rubber stopper to the test tube; invert it once or twice to mix the contents. 6 Place these test tubes at 37°C in a thermostatically controlled water bath for 30 min.

Textbook of Practical Microbiology

Note: The water level must be above the top of the milk and the bath covered with a lid to exclude the light.

QUALITY CONTROL With each test the following milk specimens are incubated as controls: A. Add 1ml methylene blue to 10ml milk that has been held at 100°C for 3 min, to inactivate its reducing system. B. Add 10 ml milk to 1 ml of tap water.

OBSERVATIONS After incubation, compare the test mixture with control A to see whether there is any decolourization in the former and with control B to see whether decolourisation is complete.

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RESULTS AND INTERPRETATION If the dye is wholly decolourised or decolourised within 5mm of the milk surface, the milk fails the test. It is considered as contaminated milk. The time for complete decolourisation need to be recorded. Because, untreated milk is often considered satisfactory if it fails to decolourize the dye in 30 minutes.

Table 85-1 List of bacteria that can be found in contaminated milk 1 2 3 4 5 6 7 8 9 10

Streptococcus lactis Streptococcus faecalis Achromobacter Clostridium perfringens Clostridium butyricum Bacillus subtilis Bacillus cereus Proteus vulgaris Staphylococcus aureus Gaffkya tetragena

KEY FACTS 1 Tuberculosis, brucellosis, streptococcal and staphylococcal infections, salmonellosis and Q fever are some of the infections transmitted by contaminated milk. 2 Methylene blue test is rapid, simple, inexpensive and most widely used test for testing the microbial contamination of the milk. 3 The time of reduction is affected by the temperature at which the milk is held before testing. The methylene blue reduction test should not be done if the atmospheric shade temperature exceeds 21°C.

VIVA 1 List common infections transmitted by contaminated milk. 2 What is the principle of methylene blue test? 3 List the routine tests used for the bacteriological examination of milk?

FURTHER READING 1 Ananthanarayanan. R, Paniker. C.K, Ananthanarayanan and Paniker’s Textbook Of Microbiology; 7th Edition, Orient Longman, pp 603 – 609, 2005. 2 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996.

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LESSON

86

Microbiology of Air

LEARNING OBJECTIVES After completing this practical you will be able to identify: 1 Bacteria present in the air. 2 Organisms present in the air by settle plate method.

INTRODUCTION Demonstration and estimation of the number of bacteria carrying particles in air may be required in certain situations. For example, in the premises where safe working depends on the bacterial content in air being kept at a very low level and premises where certain foods or pharmaceutical materials are prepared, the monitoring of density of microbial pathogens in the air is a priority. The number of bacteria in the air at any given time is dependent on the number of persons present, the amount of their body movements and the amount of disturbance of their clothing, etc. The list of bacteria commonly found in the air are summarized in the table 86-1. The infections that can be transmitted through air include wound infections, tuberculosis, anthrax, streptococcal and staphylococcal infections. Settle plate method and slit sampler method are the two methods used for routine bacteriological examination of the air. Settle plate method will be described in this chapter.

PRINCIPLE In the settle plate method, petri dishes containing an agar medium of known surface area are left open for a measured period of time. Large bacteria carrying dust particles settle on to the medium. The plates are incubated and a count of the colonies formed shows the number of settled particles that contained bacteria capable of growth on the medium.

REQUIREMENTS I Equipments Incubator and plate microscope.

II Reagents and lab wares These include Petri dishes containing nutrient agar media for the growth of organisms.

PROCEDURE 1 Prepare nutrient agar media, pour it into plates and dry off any surface moisture. 2 Remove cover of the Petri dish in its chosen position for the measured period of time, and then replace its lid. 3 Incubate the plates aerobically for 24 hours at 37°C. 4 Count the colonies, preferably with the use of a plate microscope.

QUALITY CONTROL The Petri dish agar plates should remain open for a specified and adequate time.

OBSERVATIONS Observe the colonies grown on the medium after incubation.

RESULTS AND INTERPRETATION Growth rate on the medium in a given time indicates the bacterial load in a given area. Table86-1 List of bacteria commonly found in air 1 2 3 4 5 6 7

Staphylococcus aureus Streptococcus pyogenes Mycobacterium tuberculosis Pseudomonas aeruginosa Bacillus anthracis Bacillus subtilis Proteus vulgaris

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KEY FACTS 1 The number of bacteria in the air at any given time is dependent on the number of persons present, the amount of their body movements and the amount of disturbance of their clothing, etc. 2 Settle plate method and slit sampler method are the two methods used for routine bacteriological examination of the air.

VIVA 1 List the infections transmitted by air. 2 What are the methods used for the detection of microorganisms in air?

FURTHER READINGS 1 Ananthanarayanan. R, Paniker. C.K, Ananthanarayanan and Paniker’s Textbook Of Microbiology; 7th Edition, Orient Longman, pp 603 – 609, 2005. 2 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996.

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UNIT

XIII Animal Experiments

Lesson 87

Intravenous Inoculation into Mice Tail Vein

Lesson 88 Collection of Blood from the Marginal Ear Vein of Rabbit Lesson 89 Animals and their uses in the Laboratory

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LESSON

87

Intravenous Inoculation into Mice Tail Vein

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to:

Procedure for loading the syringe for injection

1 Inoculate the appropriate culture suspension into the tail vein of mouse.

1 Open the culture tube near the flame. 2 Fill the syringe with the culture to be injected. Note: The air is to be expelled by pushing the needle into the cotton inserted into a sterile tube and the volume to be adjusted 3 The needle should be recapped carefully and placed ready for use.

INTRODUCTION Mouse is one of the most commonly used lab animal for different animal experiments. The suckling mouse is used for isolation and cultivation of viruses like Coxsackie A and B viruses and arboviruses, and also for testing enterotoxins. The adult mouse is used for isolation and cultivation of organisms such as Mycobacterium leprae, Toxoplasma gondii, Cryptosporidium parvum, etc. (Box 87-1).

PRINCIPLE Mouse is a very suitable laboratory animal for different types of pathogens. After inoculation of the animal with appropriate specimens, the changes can then be observed in the inoculated mouse and result interpreted.

Animal preparation for injection 1 Remove the selected animal from the cage. 2 Keep the animal on a rough surface such as over a mouse cage or a wire. 3 Lift the mouse by holding the tail halfway. 4 Insert the tail through the mesh of a wire basket and hold the animal in such a way that the animal is inside the wire mesh and tail outside. 5 Wipe clean the tail with spirit or antiseptic soaked cotton. 6 Apply little xylol over the area of the vein, and allow it to act for a minute so that the vein will be prominent.

Injection of material REQUIREMENTS I Equipments Mice cage. II Reagents and lab wares Discarding jar, Bunsen flame, 1 ml syringe, 26G needle, culture tube, cotton, colour dye and other standard laboratory glassware. Anaesthetic agent, and antiseptic. III Specimen Laboratory bred mouse and culture material to be tested.

1 Hold the syringe and needle in such a way that animal should be horizontal to the tail and vein. 2 Then insert the needle into the vein midway between base and tip of tail. Note: It should be ensured that the needle is inside the vein by withdrawing some blood. 3 Inject carefully the material (approx. 0.1 ml and 0.2 ml) into the tail vein. 4 Remove the needle, then place plain sterile cotton over the prick and hold it firmly until bleeding stops. 5 Keep back the animal in the cage after labeling with colored dye.

Textbook of Practical Microbiology

6 Clean the table after performing the experiment and then dispose off the needle into a discarding jar.

QUALITY CONTROL 1 The mouse has to be marked well before use for appropriate selection of animal. 2 The mouse, before inoculation should be tested for any infection or other physiological changes.

OBSERVATIONS The inoculated mouse should be observed and handled with care every day and any change in its health condition should be monitored.

RESULTS AND INTERPRETATION Depending on the type and size of inoculation, the animal will show changes and typical characteristic health manifestations. These are to be noted and interpreted.

BOX 87-1 USES OF MICE IN LABORATORY Suckling mice Less than 48 hours old weighing 0.5 to 1.0 gm. 1 For testing enterotoxins of enteropathogenic bacteria. 2 For isolation and cultivation of coxsackie A and B virus, and arbovirus. Adult mice 1 For isolation and cultivation of (a) (b) (c) (d)

Mycobacterium leprae – foot pad. Toxoplasma gondii – intraperitoneal. Cryptosporidium species - intestinal. Plasmodium berghei - tail.

2 For isolation of the causative agents namely (a) (b) (c) (d) (e) (f) (g)

Borrelia recurrentis. Francisella tularensis. Rickettsia species. Trypanosoma brucei. Chlamydia psittaci. Rabies virus. Histoplasma capsulatum.

3 For pathogenicity and virulence testing of (a) (b) (c) (d) (e) (f) (g) (h)

Streptococcus pneumoniae. Listeria monocytogenes. Bacillus anthracis. Bordetella pertussis. Nocardia asteroides. Leishmania spices. Herpes simplex virus. Cryptococcus neoformans.

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Intravenous Inoculation into Mice Tail Vein

KEY FACTS 1 The mouse to be inoculated should be first checked thoroughly for proper health conditions. 2 The area for inoculation should be disinfected before inoculation.

VIVA 1 What are the common uses of mice as laboratory animal? 2 What care should be taken during handling the mouse? Ans. The mouse should be handled only with gloved hands. It should be handled gently and the handler should thoroughly disinfect his hands after the inoculation. 3 What post inoculation care should be given for mice? Ans. After inoculation, the mouse should be kept in an individual cage with proper labeling. The animal should be fed and watered well. Its health should be monitored regularly.

FURTHER READINGS 1 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 2 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996.

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LESSON

88

Collection of Blood from the Marginal Ear Vein of Rabbit

LEARNING OBJECTIVES

PROCEDURE

After completing this practical you will be able to:

1 Restrain the rabbit by gently wrapping it in a blanket. 2 Shave the hair from the rear margin of an ear. Smear the skin over the marginal ear vein with petroleum jelly to delay clotting. 3 Grasp the ear near its base so that the venous return is impeded and the vein raised. 4 With a scalpel blade make a diagonal cut across the vein. Note: The cut should penetrate the skin and vein but not be so deep as to sever the vein. The ear should not be released at this time. 5 Blood will then start to flow. When the flow begins to recede, wiping the cut with cotton will restart it. 6 Collect the blood in a clean sterile test tube. 7 After collection release pressure on the vein, cover the cut with a small piece of cotton and press firmly. 8 Clean the ear with a sterile swab and return the rabbit to its cage.

1 Collect the blood from the ear marginal vein of rabbit.

INTRODUCTION Rabbit is one of the most commonly used lab animal for different experiments. Infant rabbits are used for the propagation of Vibrio cholerae, and for testing of enterotoxins of enteropathogenic pathogens. Adult rabbits are used for the isolation of Listeria monocytogenes, Streptococcus pneumoniae, etc. Uses of rabbits are summarized in the box 88-1.

PRINCIPLE QUALITY CONTROL

Rabbit is widely employed for raising and production of antibodies against specific antigens. Antigen is injected into rabbit and antibodies are produced. These antibodies can be obtained by drawing the blood from marginal ear vein of the rabbit. Blood samples can also be collected from central artery of the ear or directly from the heart by cardiac puncture.

1 The rabbit has to be marked well before use for ease in identification. 2 The rabbit, before inoculation should be tested for any infection or other physiological changes.

REQUIREMENTS

OBSERVATIONS

I Laboratory wares Jar, Bunsen flame, test tube, cotton, colour dye, and sterile scalpel.

The inoculated rabbit should be observed and handled with care every day and any change in its health condition should be monitored.

II Reagents Antiseptic, and petroleum jelly.

RESULTS AND INTERPRETATION

III Specimen Laboratory bred adult healthy rabbit weighing 1.5- 3 Kg.

Repeated samples of blood upto 50 ml can be collected at 2-4 week intervals.

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BOX 88-1 USE OF RABBITS IN LABORATORY Infant rabbit 1. Animal model for Vibrio cholerae. 2. In enteropathogenic organisms, testing of enterotoxins can be done. Adult rabbit 1. For testing virulence and pathogenicity of (a) Listeria monocytogenes. (b) Streptococcus pneumoniae. (c) Nocardia spp. (d) Mycobacterium tuberculosis. (e) Herpes simplex virus. (f) Candida albicans. 2. Other uses (a) Differentiating M. tuberculosis and M. bovis. (b) Sereny’s test. (c) Rabbit ileal loop test (enterotoxin testing). (d) Maintenance of Treponema pallidum. (e) Increasing virulence of rabies virus and vaccinia virus. (f) For collecting blood for antistreptolysin O test. (g) To raise antibodies to antigen. (h) For serum as source of complement for cytotoxic assays.

KEY FACTS 1 After cutting the ear and vein, the ear should not be released immediately. 2 Sudden movements or noise should not be made during the procedure. 3 Blood samples can also be collected from central artery of the ear or directly from the heart by cardiac puncture.

VIVA 1 What are the other routes for the collection of blood from rabbit?

FURTHER READINGS 1 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 2 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996.

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LESSON

89

Animals and their uses in the Laboratory

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know the commonly used laboratory animals (mice, rabbit, guinea pig and other animals) for animal experiments. 2 Know the uses of these animals in the laboratory.

After inoculation following care of the animals should be taken: 1 Physical properties of the animal should be monitored regularly. 2 Health of the animal should be checked regularly. 3 Make sure that the animal is taking normal amount of food and water as before to inoculation. 4 Take care of animal to prevent it from other infections. 5 Test results should be monitored and recorded regularly.

INTRODUCTION Laboratory animals have been used for ages, in laboratories for different purposes. These animals include mice, rabbits, guinea pigs, hamsters, monkeys, etc. They are used for a wide range of applications and form an integral part of research (Box 89-1). Animals such as these should be properly taken care of, and should be in accordance to the governing rule for animal usage in laboratory. Different animals and their usage are listed in the table 89-1.

PRINCIPLE The principle behind using of laboratory animals are that when an experimental material is inoculated into a laboratory animal, it shows pathological changes which can be later studied. The routes of inoculation in various laboratory animals include subcutaneous, intradermal, intraperitoneal, intrathecal, intravenous, intramuscular, intracerebral, etc. These different routes are selected depending on the nature of the inoculum and it is important because different organisms have different tissue tropism. The information that are to be recorded before and after inoculation of a lab animal include the number of the animal, age of the animal, physiological condition of the animal, nature of the specimen, date of inoculation of the specimen and route of inoculation. It should also include the changes noted in animals on a daily basis and the date of death of the animal if the inoculated specimen proves fatal.

The animal if dies due to the effect of the specimen inoculated, then the animal should be disinfected and incinerated with proper care.

REQUIREMENTS I Equipments Wire cage. II Reagents and animals Syringe with needle, wire mesh, hood with Bunsen flame, personnel protective equipment, masks, spirit/antiseptic and soaked cotton. Animals such as mouse (Fig. 89-1), rabbit (Fig. 89-2) and guinea pig (Fig. 89-3). III Specimen Specimen to be inoculated may be toxins including endotoxin, exotoxin and enterotoxin; bacterial, fungal or viral culture; pyrogen, etc.

PROCEDURE 1 Different routes of inoculation are followed for different types of inoculum and lab animals. 2 Commonly used routes of inoculation include intravenous, intraperitoneal, intra-abdominal, intracranial, intra-nasal, intramuscular, intradermal, subcutaneous, etc.

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3 Whichever are the lab animal used and whichever route of inoculation be the case the following general procedure can be adopted. a The animal be it mouse, rabbit or guinea pig, they are properly labeled first before use. b In case of mice, the mouse is held tightly in one hand with the tail running between two fingers. c In case of rabbit, the animal is kept inside a rabbit holding box. 4 If box is unavailable, keep the rabbit on a towel and wrap it completely leaving the head outside in such a way that the animal does not move, and is comfortable. 5 Guinea pigs, hamsters can also be held using a cloth wrapped around them. 6 The syringe is loaded with the appropriate material and gently introduced into the marked area of inoculation. 7 The area of prick can be later wiped with dry cotton and after monitoring the animal for any adverse reaction, it can be left in its cage.

FIGURE 89-1 Mouse.

QUALITY CONTROL 1 The laboratory animal’s health and physical condition should be checked before using them. 2 The animal should be labeled properly before and after inoculation.

FIGURE 89-2 Rabbit.

OBSERVATIONS The animal is marked or tagged with all necessary information like the number, date of inoculation, route of administration, type of inoculation etc. and kept in a place away from other animals.

RESULTS AND INTERPRETATION Depending upon the changes that the animal shows, the results of the test shall be interpreted.

FIGURE 89-3 Guinea pig.

BOX 89-1 USES OF LABORATORY ANIMALS To test endotoxin and exotoxin production by various bacteria. To study immune response to an antigen. To produce monoclonal and polyclonal antibodies against select antigens. To study hypersensitivity by producing reactions due to introduction of allergens. To produce complements. To use the blood/serum of animals. To test and producte vaccines. To test for pyrogens. To study changes in the condition of animals by inoculating test substances.

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Table 89-1 Laboratory animals and their usage Monkeys Monkeys are primates and are extensively used in laboratories. Since they are genetically evolved animals, they are considered very suitable for lab testing. Uses 1 To raise antiserum. 2 For RBC for serological tests. 3 For testing virulence of Entamoeba histolytica. 4 Testing pathogenicity of polio virus. 5 Testing hemagglutinating property of measles virus. 6 Laboratory cultivation of hepatitis A virus and Plasmodium falciparum. 7 Animal model for influenza virus. Hamsters Uses Infant hamsters 1 To test tumorigenicity of adeno viruses. Adult hamster 1 For pathogenicity testing of (a) Leishmania donovani. (b) Entamoeba histolytica. (c) Bacillus anthracis. 2 For isolation of (a) Leptospira species. (b) Leishmania species. (c) Toxoplasma gondii. 3 Other uses (a) Maintenance of virulence of Entamoeba histolytica. (b) Drug efficiency testing against Leishmania donovani. Guinea pig Uses 1 For the isolation of (a) Mycobacterium tuberculosis. (b) Mycobacterium bovis. (c) Yersinia pestis. (d) Leptospira icterohaemorrhagiae. (e) Brucella abortus. (f) Chlamydia species. (g) Toxoplasma gondii. 2 For toxigenicity testing of (a) Corynebacterium diphtheriae. (b) Clostridium perfringens. (c) Clostridium tetani. (d) Clostridium botulinum. 3 Pathogenicity testing of (a) Listeria monocytogenes. (b) Bacillus anthracis. (c) Clostridium perfringens. (d) Entamoeba histolytica. (e) Cryptococcus neoformans. 4 Other uses (a) For the preparation of complement. (b) For the evaluation of bronchodilator compounds. (c) Pathogenicity testing of M. tuberculosis (spleen enlargement, lesions in spleen and liver). (d) Pyrogen testing. (e) Sereny’s test.

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KEY FACTS 1 Animals should be properly taken care of, and should be in accordance to the governing rule of CPSEA for animal usage in laboratory.

2 The lab animal’s health and physiological condition should be checked before and after inoculation.

VIVA 1 What are the information to be recorded about an animal and inoculation before and after inoculation of test material? 2 What are the different routes of inoculation in lab animals and why are the different routes significant?

3 How should an inoculated animal be looked after and what precautions are to be taken?

FURTHER READINGS 1 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 2 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996.

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UNIT

XIV Medical Entomology

Lesson 90 Identification of Common Insects

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LESSON

90

Identification of Common Insects

LEARNING OBJECTIVES After completing this practical you will be able to: 1 Know important identifying features of common insects.

INTRODUCTION Insects are responsible for transmitting a wide variety of infections to humans. Hence it is essential to know different insects causing infections in man and methods to identify them.

PRINCIPLE The insects can be identified by observing the killed insect under a dissecting microscope and observing their morphological features for specific identification.

2. Head has a long needle-like structure named proboscis, a pair of palpi, each situated on either side of proboscis, a pair of antennae or feelers. 3. Antennae are bushy in male. 4. Thorax is large and rounded, bearing a pair of wings dorsally and three pairs of legs ventrally. 5. Wings are characterized by a fringe of scales on the posterior border. 6. Abdomen has 10 segments.

Identifying features of Anopheles 1. All the general features of mosquitoes as mentioned above. 2. Other characteristic features are: a). Dark (blackish) or white scales on the wing veins. b). Palpi is as along as proboscis. c). When anopheles is at rest, head, thorax, abdomen are at an angle of 45° to the resting surface. d). The middle lobe of tri-lobed salivary gland is short.

Diseases transmitted Malaria

REQUIREMENTS Identifying features of Aedes I Equipments and lab wares Standard dissecting microscope, glass slides, cover slips, sterile cotton swab, glass marking pencil, etc. II Specimen Insect to be identified.

MOSQUITOES General features 1. Head is semi-globular in shape with a pair of large compound eyes.

1. All the general features of mosquitoes as mentioned above. 2. Other characteristic features are: a). Characteristic of unspotted wings and white stripes on a black body are present. b). Tip of abdomen is pointed. c). Legs are striped or banded in nature. So they are called as tiger mosquitoes. d). Palpi are short in female. e). When at rest, body shows a hunch back. These are day bitters.

Diseases transmitted 1. Dengue. 2. Dengue haemorrhagic fever.

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3 Chikungunya fever. 4 Rift valley fever. 5 Yellow fever.

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Diseases transmitted

1 Bancroftian filariasis. 2 Japanese encephalitis. 3 West Nile fever.

1 2 3 4 5 6 7 8 9 10 11 12

SAND FLY

ITCH MITE

General features

The itch mite or Sarcoptes scabies var hominis belongs to the class Arachnida.

Identifying features of Culex 1 Similar to that of Aedes aegypti, except the tip of abdomen is blunt. 2 These mosquitoes bite about midnight.

Diseases transmitted

1 2-4mm in length, smaller than mosquito. 2 Greyish brown having dark conspicuous eye and hairy body. 3 Head contains a pair of long, slender, hairy antennae and a proboscis with 16 segments. 4 Thorax bears a pair of wings and three pairs of legs. 5 Wings are upright and lanceolate shaped. The second longitudinal vein on the wing branches twice, first branching takes place in the middle of the wing. 6 Legs are longer and hairy. 7 Only females suck blood at night. 8 Males have three pairs of clappers on the last abdominal segment. 9 They do not fly, they only hop about up to 3 feet. Diseases transmitted by sand fly are summarized in the box 90-1.

HOUSE FLIES Identifying features 1 It is mouse gray in colour. 2 Body is divided in to head, thorax, and abdomen. 3 Head bears a pair of antennae, a pair of large compound eyes and a refractive proboscis, for sucking liquid food. Eyes of the male are close together and those of the female are set apart widely. 4 Thorax is marked with 2 to 4 dark longitudinal stripes, which is characteristic of the genus Musca. It bears a pair of wing and three pairs of legs. 5 Each leg is provided with a pair of pads which enable the fly to walk on highly polished surfaces. 6 Legs and body are covered with numerous short and stiff hairs called Tenent hairs which secrete a sticky substance. 7 Abdomen is segmented and shows dark and light markings.

Typhoid and paratyphoid fever. Diarrhea. Dysentery. Cholera. Gastroenteritis. Amoebiasis. Helminthic infections. Poliomyelitis. Conjunctivitis. Trachoma. Anthrax. Yaws.

Identifying features 1 2 3 4 5

It is 0.5mm in size, visible to naked eye. Body is tortoise shaped, rounded above and flattened below. Body is hairy. Has 2 pairs of legs in front and 2 pairs behind. Front legs have suckers at the end, and the hind legs end in long bristles. 6 Males are smaller than the females. 7 In females, the first 2 pairs of legs possess suckers. In males, the fourth pair of legs also have suckers. 8 Dorsal surface has backwardly pointed spines.

Diseases transmitted 1 Scabies (seven year itch), Norwegian itch of man. 2 Forage mite and house dust mite cause allergic respiratory distress.

TROMBICULID MITE It is also known as scrub typhus mite.

Identifying features 1 2 3 4

Adult mites are 1-2mm in length and are red in colour. Covered dorsally and ventrally with numerous feathered hairs. Has 4 pairs of legs. Body is constricted between the 3rd and 4th pairs of legs, giving a figure of eight appearance.

Diseases transmitted Rickettesia orientails (Orientia tsutsugamushi). Trombicula dilensis is the vector in India. Trombicula akamush is the vector in Japan.

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HARD TICK Identifying features 1 2 3 4 5 6

Sac-like body varying in size from 2mm to 10mm. Dorsoventrally flattened, and oval in shape. Cannot distinctly be separated into head, thorax, and abdomen. Contains 4 pairs of legs but no antennae. Dorsal surface is covered by a chitinous shield, called scutum. Males have scutum on the entire dorsal surface, whereas in female it is found in small portion. 7 It has anteriorly placed capitulum called head, which projects forward beyond the scutum. 8 Spiracular openings are situated behind the basal segments of the fourth pair of legs. 9 Spurs are present in coxal segments which assist in classification of ticks. 10 Pulvilli are present. 11 Only one nymphal stage is present. 12 Takes blood meal at both day and night time. 13 Remain ectoparasite on host for long time. 14 Cannot resist starvation for a long time. 15 Life Span is 3 years. Female dies after laying eggs. Diseases transmitted by hard ticks are summarized in the box 90-2.

SOFT TICK Identifying features All features of hard ticks except 1 Scutum is absent 2 Capitulum lies ventrally, mouth parts are not visible from above. 3 Spiracular openings lies behind the 3rd pair of coxal segments. 4 Coxal segments lack spurs. 5 Pulvilli are absent. 6 More than one nymphal stage is present. 7 Take blood meals only at night. 8 Remain ectoparasite only during their short feeding time. 9 They can withstand starvation for one year. 10 Life span is 16 – 21 years.

Diseases transmitted Ornithodorus species is the vector for tick borne relapsing fever caused by Borrelia duttoni. In India relapsing fever is transmitted by Ornithodorus tholozani, Ornithodorus lahorensis and Argas persicus.

RAT FLEA Identifying features 1 Light to dark brown in colour. 2 Oval in shape, 1-8mm in size.

3 Laterally compressed, which is an adaptation to move freely in hairs and feathers. 4 Contains a hard chitinous exoskeleton and backwardly directed strong bristles. 5 Demarcation into head, thorax and abdomen is not clear. 6 Head has simple eyes and short paired antennules. 7 Mouth parts are conspicuously blood sucking and pointed downwards. 8 They do not have wings, but legs are highly developed, which helps them to leap up to 10-15cms. 9 Abdomen is bulky and has 10 segments. 10 Males have coiled genitalia. 11 Females have a spermatheca at the posterior end and Genal combs are useful for species identification.

LOUSE It is also called as Pediculus humanus.

Identifying features 1 2 3 4 5

Measures 2-4mm in length. Grayish or reddish after blood meal. Wingless ectoparasite and flattened dorsoventrally. Head is narrower than thorax and has a pair of eyes. Antenna in the larval stages is short having 3 segments while in adult it has 5 segments. 6 Thorax has 6 legs, each having 5 segments. 7 Abdomen has 9 segments, it is pointed in males, the last segment has penis. In females, abdomen is bilobed. 8 The louse are of 3 different types: a Head louse – Pediculus capitus. b Body louse – Pediculus corporis. c Pubic louse – Phthirus pubis.

Diseases transmitted 1 Louse borne epidemic typhus caused by Rickettsia prowazaki. 2 Epidemic relapsing fever caused by Borrelia recurrentis. 3 Trench fever or 5 day fever caused by Bartonella quintana.

CYCLOPS Identifying features 1 2 3 4 5

Pear-shaped. Semitransparent. Varies from 0.5 to 1mm in length. Body shows anterior cephalothorax and posterior abdomen. Cephalothorax has a Pigmented eye. b Two pairs of antennae. c Three pairs of jaws (1 pair mandibles and 1 pair maxillae). d Five pairs of legs which are swimming appendages. 5th pair is vestigeal.

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6 Abdomen has 5 segments. The last segment has two feathered filaments. In females the first abdominal segment has got 2 ovisacs on both sides to carry the eggs. 7 Swims with characteristic jerky movements in water. Hence called as water fleas. They can be seen as small moving specks in water.

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Diseases transmitted 1 Dracunculiasis. Mesocyclops act as intermediate host for Dracunculosis medinensis. 2 Diphyllobothriasis. First intermediate host for Diphyllobothrium latum. 3 They also act as vectors for Gnathostoma spinigerum and Gnathostoma hispidium.

BOX 90-1 DISEASES TRANSMITTED BY SAND FLY Diseases

Species

1 Kala azar. 2 Oriental sore . 3 Sand fly fever.

Phlebotomus argentipes. Phlebotomus sergenti. Phlebotomus papatasi and Sergentomyia spp. Lutzomyia verrucarum. Lutzomyia columbiana. Lutzomyia peruensis.

4 Oroya fever . (Bartonellosis, Carrion’s disease)

BOX 90-2 DISEASES TRANSMITTED BY HARD TICKS Diseases

Species

1 Tick paralysis. 2 Colorado tick fever. 3 Kyasanur forest disease. 4 Crimean – congo hemorrhagic fever. 5 Russian spring summer encephalitis. 6 Tick borne encephalitis. 7 Osmk hemorraghic fever. 8 Lyme disease. 9 Rocky mountain Spotted fever. 10 Boutonneuse fever.

Dermacentor, Ixodes, Haemophysalis. Dermacentor andersoni. Haemophysalis spinigera. Hyalomma marginatum, Dermacentor marginatus. Ixodes persculatuis, Haemophysalis concinna. Dermacentor marginatum. Dermacentor, Ixodes. Ixodes scapularis. D. andersoni, D. variabilis. Rhipicephalus sanguineus.

VIVA 1 2 3 4

List the diseases transmitted by mosquitoes. List the identifying features of cyclops. What are the identifying features of Anopheles mosquitoes? What are the diseases transmitted by house fly?

FURTHER READINGS 1 Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. 2 Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996.

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UNIT

XV Common Viva Spots

Lesson 91 Identification of Common Viva Spots

274

LESSON

91

Identification of Common Viva Spots

LEARNING OBJECTIVES

Preparation

After completing this practical you will be able to identify the following common viva spots given in Microbiology examination:

Prepared by adding agar in nutrient broth and sterilized by autoclaving.

Modifications 1 2 3 4 5 6 7

Culture media. Culture media with growth. Biochemical reactions. Specimens of parasites. Glass wares. Instruments, and Microscopic slides.

CULTURE MEDIA

1 Semi-solid agar if concentration of agar is 0.2–0.5%. 2 Concentrated agar if agar concentration is 2–6%.

Uses 1 2 3 4

For growing of non fastidious bacteria. Determination of antibiotic sensitivity. Preparation of blood agar. Concentrated agar (3% and more) prevents swarming of bacteria such as Proteus vulgaris, Clostridium tetani, etc.

NUTRIENT AGAR BLOOD AGAR Also referred as ‘Agar’ (Fig. 91-1).

Composition Nutrient broth : Peptone water. Meat extract : 1%. Agar : 2–3%.

FIGURE 91-1 Nutrient agar.

The blood agar is an enriched medium as well as an indicator medium. The colour of the medium is red like that of blood (Fig. 91-2).

Composition Nutrient agar. Sheep blood :

5–10%.

FIGURE 91-2 Blood agar.

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Preparation Prepared by adding sterile blood to sterile nutrient agar that has been melted and cooled to 55°C. The blood agar is sterilized by autoclaving. Concentration of blood vary from 5% to 50% for special purposes. 10% is most usual concentration used. Either human or animal (sheep, rabbit, horse, etc) blood may be used.

Uses FIGURE 91-4 Mac Conkey agar.

1 For growth of most of the pathogenic bacteria. 2 Used in preparation of the potassium tellurite blood agar.

CHOCOLATE AGAR The medium is so called because it is chocolate in colour. Chocolate agar is an enriched medium (Fig. 91-3).

Composition Nutrient agar. Heated 10% sterile blood (horse, sheep).

Composition 1 2 3 4 5

Peptone Sodium taurocholate, commercial Water Agar Neutral red solution, 2 percent in 50 percent ethanol 6 Lactose 10% aqueous solution.

: : : :

20 g. 5 g. 1 litre. 20 g.

:

3.5 ml.

Preparation Dissolve peptone and taurocholate (bile salt) in water by heating. Add the agar and dissolve it in the steamer or autoclave. If necessary, clear by filtration. Adjust pH to 7.5. Add lactose and neutral red which should be well shaken before use and mix. Heat in autoclave with free steam (100°C) for 1 hr, then at 115°C for 15 min. Pour plates. The medium is sterilized by autoclaving. The medium should be a distinct reddish brown colour. If it is acid, it assumes rose-pink colour.

Uses FIGURE 91-3 Chocolate agar.

1 For growing enteric bacteria. 2 To differentiate lactose fermenter (LF) from non-lactose fermenter (NLF) colonies (Salmonella, Shigella, etc.).

Preparation It is prepared by heating 10% sterile blood in sterile nutrient agar. The agar is melted and cooled it in water bath at 75°C, blood is added and the medium continues to remain at 75°C till the blood becomes chocolate brown in colour.

LOEFFLER’S SERUM SLOPE Loeffler’s serum slope is an enriched medium. This is used for culture and isolation of Corynebacterium diphtheriae (Fig. 91-5).

Uses For growing fastidious bacteria such as Neisseria, Pneumococcus, Haemophilus, etc.

MACCONKEY AGAR MacConkey agar is a differential or indicator medium. This is a useful medium for cultivation of enteric bacteria (Fig. 91-4).

FIGURE 91-5 Loeffler’s serum slope.

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Composition 1 Sterile ox, sheep, horse serum 2 Nutrient broth 3 Glucose

Preparation : : :

300 ml. 100 ml. 1 g.

Preparation Dissolve glucose in nutrient broth and autoclave at 115°C for 20 min. Add glucose broth to the serum with sterile precautions and distribute in test tubes or in 2–5 ml amounts in sterile conditions to fill up nearly one fourth of the bottles. This medium is sterilized by inspissation.

Uses 1 Loeffler’s serum slope is used especially for cultivation of C. diphtheriae, producing luxuriant growth in 6–12 hr. 2 It is also used to show proteolytic properties, particularly of Clostridium species.

Malachite green solution: Prepare 2% solution of malachite green in sterile water with sterile precautions by dissolving the dye in incubator for 1–2 hr. Preparation of medium Mineral salt solution : Malachite green solution : Beaten egg (20 to 22 hen’s eggs depending on size) :

600 ml. 20 ml. 1 litre.

Mix the complete medium, distribute it in 5 ml volumess in sterile McCartney bottles and screw the caps tightly on. Lay the bottles horizontally in the inspissator and heat at 75°–80°C for 1 hr. This is to solidify the medium.

Uses 1 For growing Mycobacterium tuberculosis (takes 3 to 6 weeks to grow). 2 For antibiotic sensitively testing of M. tuberculosis.

LOWENSTEIN-JENSEN (LJ) MEDIUM Lowenstein-Jensen (LJ) medium is an enriched medium used for growing Mycobacterium species. The green colour of the medium is due to the presence of malachite green (Fig. 91-6).

Composition 1 Mineral salt solution Potassium dihydrogen phospphate anhydrous KH2 PO4 : 2.4 g. Magnesium sulphate, MgSO4 : 0.24 g. Magnesium citrate : 0.6 g. 2 Asparagine : 3.6 g. 3 Glycerol : 12 ml. 4 Water : 600 ml. 5 Egg. 6 Malachite green solution. Dissolve mineral salts by heating. Autoclave at 121°C for 25 min to sterilize.

FIGURE 91-6 Lowenstein Jenson’s medium.

ROBERTSON COOKED MEAT (RCM) BROTH Robertson cooked meat (RCM) broth is a selective medium used for culture of anaerobic bacteria (Fig. 91-7).

Composition 1 2 3 4 5

Cooked meat (fresh bullock heart) Peptone Sodium chloride Water Liquid from cooked meat

: : : : :

500 g. 2.5 g. 1.25 g. 500 ml. 500 ml.

Preparation Place the meat in each 1 oz bottle to a depth of one inch and cover with about 10 ml broth. The medium is sterilised by autoclaving at 121°C for 20 min. After sterilization the pH of

FIGURE 91-7 Robertson’s cooked meat medium. (RCM ) broth.

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broth over meat is 7.5. If test tubes are used the surface of the medium may be covered with a layer of sterile liquid paraffin 1cm deep but this is not essential.

meat and it is essential that basal medium to which various carbohydrates added for fermentation tests should be free from natural sugars (Fig. 91-9).

Uses

Composition

1 For growing anaerobic bacteria. 2 For preservation of stock cultures of anaerobic bacteria.

Peptone : Sodium chloride (NaCl) : Water :

10 g. 5 g. 1 L.

SABOURAUD’S DEXTROSE AGAR (SDA) Sabouraud’s dextrose agar (SDA) is used for culture of fungi. Low pH and high sugar content of this medium make it particularly selective for fungi as against bacterial contaminants (Fig. 91-8).

Composition Glucose Peptone Agar Water

: : : :

40 g. 10 g. 20 g. 1 L.

FIGURE 91-9 Peptone water.

Preparation Dissolve the ingredients in warm water, adjust pH to 7.4–7.5 and filter. Distribute as required and autoclave at 121°C for 15 min. This medium is sterilized by autoclaving.

Uses

FIGURE 91-8 Sabouraud’s dextrose agar (SDA).

1 2 3 4 5

For growing pathogenic bacteria. For making hanging drop preparation of bacteria. For preparation of sugar media. For growing bacteria for antibiotic susceptibility testing. To test the formation of indole.

Preparation Dissolve the ingredients in the steamer or autoclave. Filter through cotton gauze and adjust to pH 5.4. Dissolve in stock bottles or in tubes. Autoclave at 115°C for 15 min.

Uses

GLUCOSE BROTH Glucose added to nutrient media promotes luxuriant growth of many organisms. It also acts as reducing agent and used for shake cultures of anaerobes (Fig. 91-10).

1 For growing fungi. 2 The agar medium is suitable for the primary isolation of fungi from clinical material. 3 Fungi can take weeks to grow but yeasts grow in 24 hours to 48 hours on this medium.

PEPTONE WATER Peptone water is used as the basis for sugar fermentation media since broth may contain a small amount of sugar derived from

FIGURE 91-10 Glucose broth.

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Composition 1 2 3 4

Nutrient broth. Peptone water. Meat extract Glucose

STAPHYLOCOCCUS AUREUS ON NUTRIENT AGAR 1%. (0.5%).

Preparation Prepare 20% of glucose solution separately. Add a drop of phosphoric acid to endure pH not more than 7.0 and autoclave at 115°C for 20 mins. This sterile glucose solution is then added to the sterile basal medium.

Uses

Salient features 1 Seen as b-hemolytic colonies on blood agar with golden yellow pigment. 2 The colonies are opaque convex with a shining surface and may be pigmented white (var. albus) yellow, golden yellow or golden (var. aureus). Confluent growth appears like oil paint (Fig. 91-12). 3 Gram staining shows as Gram positive cocci in clusters. 4 S. aureus is catalase positive which differentiates it from streptococci. 5 It is coagulase positive.

1 Used for growing fastidious organisms such as Streptococcus pyogenes and Enterococci, and for their antibiotic susceptibility testing.

CULTURE MEDIA WITH GROWTH STREPTOCOCCUS PYOGENES ON BLOOD AGAR FIGURE 91-12 Staphylococcus aureus on nutrient agar.

Salient features 1 Blood agar colonies 0.5–1 mm in diameter, after 24 hrs incubation circular, discrete, semi-transparent, low convex disks, showing a-hemolysis on fresh blood agar plates (Fig. 91-11). 2 Virulent strains isolated from lesions give a matt type of colony whereas avirulent strains produce glossy colonies. 3 A mucoid colony type is also encountered and corresponds in virulence to matt type. 4 Str. pyogenes is the commonest organism causing sore throat. 5 Seen as b-hemolytic colonies in blood agar plates. 6 Gram staining shows Gram positive cocci in chains. 7 It is catalase negative. This test differentiates it from Staphylococcus.

FIGURE 91-11 Streptococcus pyogenes on blood agar.

PROTEUS SPP. ON BLOOD AGAR Salient features 1 Group of cells at the edge of developing micro colony migrate to an uninoculated area of the medium and present as swarming. Swarming is seen in blood agar plates and the growth may eventually appear either as a uniform growth / film over the whole plate. Continuous swarming or discontinuous swarming are a series of concentric circles of growth around point of inoculation (Fig. 91-13). 2 Proteus spp. are Gram negative, coccobacilli, 1–3 µm long and 0.6 µm wide. In young culture they may be filamentous as longs as (80 µm). 3 They are motile by peritrichous flagella. They possess more than one type of fimbriae. Variants are nonflagellate and non motile.

FIGURE 91-13 Proteus species on blood agar.

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4 The two common species are Proteus vulgaris and Proteus mirabilis. 5 PPA and urease tests are positive.

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279

4 They are indole positive, urease negative. 5 They cause urinary tract infections and diarrhoea.

KLEBSIELLA SPP ON MACCONKEY AGAR Salient features

Salient features 1 On nutrient agar growth of the organism shows green diffusible pigment (Fig. 91-14). 2 Ps. aeruginosa is a strict aerobe, slender, Gram negative bacillus, 1.5–3.0 µm × 0.5 µm arranged singly, in pairs or short chains. 3 It is non sporing, non capsulate, though mucoid strains may sometime occur and usually motile by one or two polar flagella. 4 It is oxidase positive. 5 It is one of the common causes of nosocomial infections.

1 Klebsiella spp produces lactose fermenting, mucoid colonies on MacConkey agar (Fig. 91-16). 2 They are Gram negative, non sporing, non motile bacilli, 1–2 µm long and 0.5–0.8 µm wide with parallel or bulging sides and slightly pointed or rounded ends. 3 Freshly isolated strains possess a well defined polysaccharide capsule. 4 They are urease positive, and indole negative. 5 They cause urinary tract infections, pneumonia, etc.

FIGURE 91-16 Klebsiella species on MacConkey agar. FIGURE 91-14 Pseudomonas aeruginosa on nutrient agar.

ESCHERICHIA COLI ON MACCONKEY AGAR Salient features 1 Escherichia coli shows lactose fermenting, non-mucoid colonies on MacConkey agar (Fig. 91-15). 2 It is an aerobe and facultative anaerobe. 3 It is a Gram negative, non capsulated bacillus measuring 1–3 µm × 0.4–0.7 µm. Most of the strains are motile by peritrichate flagella.

FIGURE 91-15 Escherichia coli on MacConkey agar.

CORYNEBACTERIUM DIPHTHERIAE ON POTASSIUM TELLURITE AGAR Salient features 1 Potassium tellurite agar is used as selective medium for C. diphtheriae (Fig. 91-17). 2 C. diphtheriae produces black coloured colonies in potassium tellurite agar.

FIGURE 91-17 Corynebacterium diphtheriae on potassium tellurite agar.

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3 S. aureus is another bacteria which produces black colonies on this medium 4 Other media used for C. diphtheriae are Loeffler’s serum slope, and blood agar containing fresh, lysed or heated blood.

MYCOBACTERIUM TUBERCULOSIS ON LÖWENSTEIN JENSEN (LJ) MEDIUM Salient features 1 M. tuberculosis takes 3–6 weeks to grow on the LJ medium on incubation at 37°C (Fig. 91-18). 2 Cultures should not be discarded as negative until they have been observed for 12 weeks. 3 The colonies are identified by their rough, tough and buff coloured eugonic growth.

BIOCHEMICAL REACTIONS CARBOHYDRATE FERMENTATION TESTS Salient features 1 It is used to determine the ability of an organism to ferment a particular carbohydrate to produce acid or acid and gas (Fig. 91-20). 2 A large variety of sugars are used as follws: Pentoses Hexoses

: :

Disaccharides

:

Trisaccharides : Polysaccharides : Sugar alcohols : Glycosides

FIGURE 91-18 Mycobacterium tuberculosis on Lowenstein- Jensen’s medium.

CANDIDA ALBICANS ON SABOURAUD’S DEXTROSE AGAR (SDA) Salient features 1 Candida albicans species grow well on SDA at 25–37°C (Fig. 91-19). 2 Cream coloured, smooth pasty colonies appear in 1–2 days. 3 Lacto-phenol cotton blue preparation and Gram stained smears showing budding yeast cells and pseudohyphae. 4 C. albicans can be differentiated from other species by germ tube test, sugar fermentation and sugar assimilation reactions

:

3 A suitable indicator that will change colour only as a result of formation of acids during fermentation of sugar is used in the test. A small inverted tube (Durham’s fermentation tube) is placed in each culture tube to detect gas. Andrade’s indicator is the indicator used. Pink colour of the solution is due to acid production by fermentation of carbohydrates. Air bubble is formed in the Durham’s tube if gas is produced. 4 The test is done by inoculating a drop or loopful of the culture. The inoculated tubes are incubated, aerobically at 37°C for required period, and colour change and gas formation noticed. Fermentation reactions are observed usually after a period of 24 hours of incubation. 5 These tests are used to identify some Gram negative bacilli such as E. coli, Klebsiella species, Salmonella species, Proteus species etc.

+ FIGURE 91-19 Candida albicans on Sabourauds dextrose agar.

Arabinose, xylose, rhamnose. Glucose, fructose, mannose, sorbose, galactose. Sucrose, maltose, lactose, trehalose, cellobiose. Raffinose. Starch, insulin, dextrin, glycogen. Glycerol, erythritol, adonitol, mannitol, dulcitol, sorbitol, inositol. Salicin, aesculin.

+

+

–

FIGURE 91-20 Carbohydrate fermentation test.

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GLUCOSE WITH DURHAM’S TUBE Salient features 1 It is used to determine the ability of an organism to ferment a particular carbohydrate to produce acid or acid and gas (Fig. 91-21). 2 All members of the family Enterobacteriaceae are glucose fermenters with or without gas production. 3 This test is used to identify some Gram negative bacilli such as E. coli, Klebsiella species, Salmonella species, Proteus species, etc.

–

+

281

4 Examples of indole positive bacteria are E. coli, Proteus vulgaris, Edwardsiella species, etc. 5 Kovac’s reagent consists of amyl iosoamyl alcohol, 150ml; p-dimethyl – amino benzaldehyde; 10g and conc. HCl; 50ml.

UREASE TEST Salient features 1 This test determines the ability of an organism to produce an enzyme urease which splits urea to ammonia (Fig. 91-23). 2 The occurrence of this enzyme, urease can be tested for alkali (NH3) production by means of suitable pH indicator. 3 Christensen’s urease medium is an example of the medium which contains phenol red as an indicator. Ammonia produced from urea makes the medium alkaline and phenol red changes to pink / red in colour. 4 Development of pink colour indicates positive test while the persistence of the pale yellow colour indicates the negative test. 5 Examples of urease producing bacteria are Klebsiella species, Proteus species, Yersinia enterocolitica, Helicobacter pylori,etc.

FIGURE 91-21 Glucose with Durham’s tube.

INDOLE TEST Salient features 1 The indole test demonstrates the ability of certain bacteria to decompose the amino acid tryptophan to indole which accumulates in the medium. The production of indole is then tested for by a colorimetric reaction with p-dimethyl amino benzaldehyde (Fig. 91-22). 2 The test is performed by inoculating the peptone water medium with bacterium to be tested and incubated for 24-48 hours at 37°C. Then 0.5ml Kovac’s reagent is added to the medium and shaked gently. 3 Formation of a red coloured ring near the surface of medium indicates positive test. The presence of yellow coloured ring near surface of medium indicates negative test.

+ FIGURE 91 -22 Indole test.

NEGATIVE POSITIVE

FIGURE 91-23 Urease test.

CITRATE UTILIZATION TEST Salient features 1 This is a test for the ability of an organism to utilize citrate as the sole carbon and energy source for growth and an ammonium salt as the sole source of nitrogen (Fig. 91-24). 2 Koser’s liquid citrate medium or Simmon’s citrate agar may be used. Bromothymol blue is the indicator in Simmon’s citrate medium. 3 Positive Koser’s citrate medium is indicated by the formation of a turbidity i.e. growth. No turbidity indicates negative growth. 4 In Simmon’s citrate medium positive test is indicated by formation of blue colour and streak of growth. Original green colour and no growth indicates negative growth. 5 Examples of citrate positive bacteria are Klebsiella species, Salmonella species except Salmonella Typhi, Citrobacter species, etc.

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NEGATIVE

POSITIVE

FIGURE 91-24 Citrate utilization test.

FIGURE 91-26 Triple sugar iron agar.

PHENYL PYRUVIC ACID TEST (PPA) Salient features 1 It is used to determine the ability of an organism to deaminate phenyl pyruvic acid (Fig. 91-25). 2 Positive PPA test is indicated by a green colour and negative PPA test is indicated by no colour change. Examples of PPA positive bacteria are Proteus species, Providencia species, and Morganella species.

4 Yellow colour formation occurs with fermentation of carbohydrates, while bubbles in butt show the formation of gas during fermentation process. 5 When H2S is produced by the bacteria, the medium shows blackish discolouration. 6 Combinations of TSI reactions: K/A (red/yellow) A/A (yellow/yellow)

: :

Glucose only fermented. Glucose, and lactose or sucrose fermented or both fermented. K/K (red / red) : Neither glucose, lactose nor sucrose fermented. Note: K – Alkaline, A- Acidic.

SPECIMENS OF PARASITE ASCARIS LUMBRICOIDES ADULT WORM Salient features NEGATIVE

POSITIVE

FIGURE 91-25 Phenyl pyruvic acid test.

1 Tail end of male worm is curved ventrally in form of a hook, while in female worm, it is conical and straight (Fig. 91-27). 2 Route of infection of the organism is by ingestion of the food or water contaminated with eggs. 3 Embryonated egg is the infective form of the parasite.

TRIPLE SUGAR IRON (TSI) AGAR Salient features 1 TSI agar is used to determine the ability of the bacterium to break down specific carbohydrates incorporated in a growth medium, with or without the production of gas, along with the production of hydrogen sulphide (Fig. 91-26). 2 Three carbohydrates i.e., glucose, lactose and sucrose are present in the TSI agar. It also contains ferric salts for detection of H2S production. 3 The medium in a test tube has a butt and slant. Phenol red is the indicator.

FIGURE 91 -27 Ascaris lumbricoides adult worm.

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HYDATID CYST

BIJOU BOTTLE

Salient features

Sterilised by hot air oven or autoclave (Fig. 91-30).

1 Caused by Echinococcus granulosus. 2 Cyst wall consists of two layers (i.e.) ectocyst and endocyst (Fig. 91-28). 3 Hydatid fluid which is present in cyst is used as antigen in immunodiagnostic tests. 4 Man is the intermediate host and dog is the definitive host. 5 Infective agent is the egg, which is present in dog’s faces. 6 Infection is acquired by ingestion of the food or water contaminated eggs.

Uses

283

1 Specimen collection like CSF, blood, urine, ascitic fluid, etc. 2 For preparation of media e.g. LJ medium, Loeffers, serum slope, etc. 3 For preparation of urease medium.

FIGURE 91-30 Bijou bottle. FIGURE 91-28 Hydatid cyst.

GLASS SYRINGE GLASS WARES

Glass syringes (Fig. 91-31) are sterilised by hot air oven.

Uses UNIVERSAL CONTAINER Universal containers (Fig. 91-29) are sterilised by autoclave .

Uses

1 For collection of blood by venepuncture. 2 To collect body fluids, pus, etc. 3 To collect blood from animals (sheep, rabbit) which may be used for preparation of blood agar. 4 For injecting medicine to patients.

1 Specimen collection. 2 For measuring and mixing purposes.

FIGURE 91-31 Glass syringe.

TUBERCULIN SYRINGE

FIGURE 91-29 Universal containers.

These are of two types: Glass or plastic syringe, and graduated 1ml syringe (Fig. 91-32). a) Glass syringes are sterilized by hot air oven. b) Plastic syringes are sterilized by gamma radiations.

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Identification of Common Viva Spots

FIGURE 91-32 Tuberculin syringe.

FIGURE 91-34 Graduated pipette.

Uses

PASTEUR PIPETTE

1 2 3 4 5 6

Pasteir pipettes (Fig. 91-35) are sterilised by hot air oven.

Lepromin test. Tuberculin test. BCG vaccination. Insulin injection. Tetanus toxoid injection. To give intradermal and other sub cutaneous injections; and to inject small amount of test material into animals.

Uses 1 Used for delivering solutions or reagent in test tubes, containers etc., during various procedures. 2 For mixing the constituents for reactions. Example: RBC s coated with specific antigen to perform IHA test for detection of antibodies.

PETRI DISH Petri dishes (Fig. 91-33) are sterilised by hot air oven.

Uses 1 For preparation of culture media such as nutrient agar, blood agar, MacConkey agar, etc. 2 For counting colonies as in pour plate method. FIGURE 91-35 Pasteur pipette.

SWAB TUBE Swab tubes (Fig. 91-36) are sterilized by hot air oven. Unplugged swab tubes are contaminated. Presterilized disposable swabs are commercially prepared. These are sterilized by gamma radiations.

FIGURE 91-33 Petri dish.

GRADUATED PIPETTE These are of 2 types: measuring pipette (Fig. 91-34) and delivery pipette. Sterilised by hot air oven.

Uses 1 For measuring quantity of fluid used in serological tests or other tests. 2 For delivering the exact required volume.

FIGURE 91-36 Swab tube.

Textbook of Practical Microbiology

Uses 1 For collection of specimens from various sites e.g. throat, cervix, local lesions, etc. 2 For streaking in case of lawn culture of bacterial growth as in antibiotic sensitivity testing

NIH SWAB

285

HOT AIR OVEN Salient features 1 Hot air oven (Fig. 91-39) uses the principle of dry heat sterilization (holding temperature at 160°C for 60 min) method. 2 It is used for sterilization of glass wares such as glass syringes, Petri dish, flasks, pipettes, test tubes, etc.

This is a swab used for collection of perianal scrapings.

Uses 1 Specially used for collection of specimen from perianal region for demonstration of eggs of Enterobius vermicularis (Fig. 91-37).

FIGURE 91-37 Egg of Enterobius vermicularis, x 400.

INSTRUMENTS INCUBATOR Salient features 1 Incubators (Fig. 91-38) are used for incubating culture plates and liquid media at specified temperature for growth of micoorganisms such as bacteria, fungi, amoebae, etc. 2 Optimum temperature for growing bacteria is 37°C. 3 It contains a thermometer by which periodically temperature can be monitored.

FIGURE 91-38 Incubator.

FIGURE 91-39 Hot air oven.

AUTOCLAVE Salient features 1 Autoclave (Fig. 91-40) uses the principle of moist heat sterilization (121°C at 15 lbs pressure for period of 15 minutes) method. 2 It is used for sterilization of culture media, rubber material, gloves, gowns, dressing, test tubes, etc.

FIGURE 91-40 Autoclave.

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Identification of Common Viva Spots

MICROSCOPY SLIDES

NEISSERIA GONORRHOEAE Salient features

GRAM POSITIVE COCCI Salient features 1 In Gram stained smear, the Gram positive cocci appear violet in colour (Fig. 91-41). 2 Gram positive cocci occur either in pairs, chains or clusters.

1 In Gram stained smear, Neisseria gonorrhoea appear as pink coloured Gram negative diplococci with adjacent sides concave typically kidney shaped (Fig. 91-43). 2 Found predominantly within the polymorphonuclear cells, but some may be seen outside.

Examples S. aureus Streptococcus pneumoniae Micrococci Sarcina species Streptococcus pyogenes

: : : : :

Arranged in clusters. Arranged in pairs. Arranged in tetrads. Arranged in groups of eight. Arranged in chains.

FIGURE 91-43 Neisseria gonorrhoeaein a Gram’s stained smear, x 1000.

GRAM NEGATIVE BACILLI Salient features FIGURE 91-41 Gram positive cocci, x 1000.

STREPTOCOCCUS PNEUMONIAE Salient features 1 In Gram stained smear, Str. pneumoniae appear as Gram positive lanceolate shaped cocci arranged in pairs, surrounded by capsule (Fig. 91-42). 2 Capsule is seen as a clear halo around the diplococci. 3 Capsule of Str. pneumoniae can be typically demonstrated by negative staining with India ink.

FIGURE 91-42 Sterptococcus pneumoniae in a Gram’s stained smear, x 1000.

1 In Gram stained smear, the Gram negative bacilli appear red in colour (Fig. 91-44). 2 They usually do not have any typical arrangement 3 In capsulated organisms, capsule can be seen as unstained structure surrounding the cell.

Examples E. coli, Klebsiella species, Proteus species, etc.

FIGURE 91-44 Gram negative bacilliin a Gram’s stained smear, x 1000.

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HAEMOPHILUS INFLUENZAE Salient features 1 Gram stained smear shows pink coloured Gram negative pleomorphic bacilli of Haemophilus influenzae (Fig. 91-45). 2 Coccobacillary forms of the bacteria can also be seen.

FIGURE 91-47 Bacillus anthracis in a Gram’s stained smear, x 1000.

CLOSTRIDIUM PERFRINGENS Salient features

FIGURE 91-45 Haemophilus influenzaein a Gram’s stained smear, x 1000.

1 Gram stained films of necrotic muscle tissue shows Gram positive bacillus of 4-6 µm x 1 µm with straight, parallel sides and rounded or truncated ends without spores occurring singly or in chains (Fig. 91-48). 2 Pus cells are absent or scanty.

VIBRIO CHOLERAE Salient features 1 Gram stained smear shows pink coloured comma shaped, curved Gram negative bacilli with rounded or slightly pointed ends (Fig. 91-46). 2 Vibrios are seen arranged on parallel rows fish in stream appearance.

FIGURE 91-48 Clostridium perfringens in a Gram’s stained smear, x 1000.

TREPONEMA PALLIDUM Salient features FIGURE 91-46 Vibrio cholerae in a Gram’s stained smear, x 1000.

BACILLUS ANTHRACIS

1 Treponema pallidum in a smear stained by Levaditi stain appear as thin delicate spirochaetes of 10 µm long with about 10 regular spirals which are sharp and angular at regular intervals of about 1 µm (Fig. 91-49). 2 Seen as black spirals against yellowish brown background

Salient features 1 In a Gram stained smear, Bacillus anthracis appears as short chains of thick purple coloured Gram positive rods arranged end to end surrounded by capsule (Fig. 91-47). 2 Ends of the bacilli are truncated, often concave and somewhat swollen giving the chain of bacilli a bamboo stick appearance.

FIGURE 91-49 Triponema pallidum in a Levaditi stained smear, x 1000.

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Identification of Common Viva Spots

ALBERT STAINING Salient features 1 It is a special stain for demonstrating metachromatic granules of C. diphtheriae (Fig. 91-50). 2 The bacteria appears as green coloured bacilli with bluish black metachromatic granules, arranged in Chinese letter pattern (V or L pattern). 3 Green colour of the bacilli is due to malachite green and bluish black granules due to toluidine blue reagents of Albert’s stain. 4 C. diphtheriae causes diphtheria. 5 Potassium tellurite blood agar is the selective medium for C. diphtheriae. The bacteria forms black colour on this medium after 36-48 hrs of incubation at 37°C. 6 Loeffler’s serum slope is an egg based medium , on which colony of C. diphtheriae can be seen as early as 6-8 hours after incubation.

FIGURE 91-51 Ziehl Neelsen staining for Mycobacterium tuberculosis, x 1000.

ZIEHL - NEELSEN STAINING FOR MYCOBACTERIUM LEPRAE Salient features 1 It is a special stain for demonstrating acid fastness of Mycobacterium leprae (Fig. 91-52). 2 5% sulphuric acid is used for ZN stain. M. leprae resist decolourisation with 5% sulphuric acid. 3 M. leprae is only acid fast but not alcohol fast. 4 M. leprae are seen as pink coloured acid fast bacilli arranged as parallel rows of bacilli giving a ‘cigar bundle’ or ‘globi’ appearance against a blue background. 5 M. leprae causes leprosy. 6 No artificial cell-free media is available for growth of M. leprae, hence it does not satisfy Koch’s postulates for a pathogenic bacteria. 7 It can be grown in foot pads of 9 banded armadillo.

FIGURE 91-50 Alberts stained smear of Corynebacterium diphtheriae, x 1000.

ZIEHL - NEELSEN STAINING FOR MYCOBACTERIUM TUBERCULOSIS Salient features 1 It is a special stain for demonstrating acid fastness of M. tuberculosis (Fig. 91-51). 2 20% sulphuric acid is used for ZN stain. M. tuberculosis resist decolourisation with 20% sulphuric acid. 3 M. tuberculosis is both acid and alcohol fast. 4 M. tuberculosis appear as approx. 1-3 µm size slender pink coloured acid fast bacilli with curved ends against a blue background. 5 M. tuberculosis causes tuberculosis. 6 Löwenstein Jensen (LJ) medium is the most frequently used medium to grow M. tuberculosis. 7 It takes 3-6 weeks for M. tuberculosis to grow in this medium.

FIGURE 91-52 Ziehl Neelsen staining for Mycobacterium leprae, x 1000.

NEGRI BODIES Salient features 1 Negri bodies are 3 - 27 µm size, intra cytoplasmic inclusion body which appear as round , oval, purplish pink structures with characteristic basophilic inner granules (Fig. 91-53). 2 Negri bodies are usually found within the nerve cells of the hippocampus and cerebellar region. 3 These are usually demonstrated in the impression smears of

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the rabid dog brain stained by Seller’s technique (basic fuschin and methylene blue in methanol). 4 Negri bodies can also be demonstrated by Mann’s and Giemsa stain. 5 Presence of basophilic inner granules within the Negri bodies helps to differentiate canine distemper which lacks it.

FIGURE 91- 55 Molluscum bodies, x 1000.

PLASMODIUM VIVAX RING STAGE Salient features

FIGURE 91- 53 Negri bodies, x 1000.

MULTINUCLEATE GIANT CELLS: MEASLES Salient features

1 Plasmodium vivax is the causative agent of vivax malaria (Fig. 91-56). 2 Female Anopheles mosquito transmits the disease and is the definitive host 3 Man is the intermediate host. 4 It is a Giemsa stained thin blood smear, the ring stage appear as a blue stained ring of cytoplasm with a red chromatin dot. The cytoplasm portion of ring opposite the nucleus is thickened. 5 Young erythrocytes are predominantly infected and they are enlarged.

1 These multinucleate giant cells are produced by measles virus in Hela cell lines in tissue culture (Fig. 91-54). 2 These giant cells show multinucleate syncytium formation, with numerous acidophilic nuclear and cytoplasmic inclusions in Giemsa stained smears.

FIGURE 91- 56 Plasmodium vivax ring stage in a Giemsa stained blood smear, x (1000.

PLASMODIUM FALCIPARUM RING STAGE FIGURE 91- 54 Multinucleate giant cells-measles, x 1000.

MOLLUSCUM BODIES Salient features 1 Section of the lesion shows large (20 µm to 30 µm) sized eosinophilic hyaline inclusion bodies with nuclei displaced to the margin (Fig. 91-55). 2 Molluscum bodies are composed of large nucleus of virus particles, embedded in protein matrix.

Salient features 1 In a Giemsa stained thin blood smear, ring stage appears as blue ring of cytoplasm surrounding a central vacuole with red chromatin dot at centre. 2 P. falciparum is the causative agent of falciparum malaria and causes cerebral malaria (Fig. 91-57). 3 Multiple ring forms are found within a single RBC, along with accole forms. 4 All ages of erythrocytes are infected. Ring stages are found within both normal sized young and older RBCs. 5 The erythrocyte which is invaded by the parasite is not enlarged 6 P. falciparum shows frequent drug resistance to chloroquine.

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Identification of Common Viva Spots

MALE FEMALE

FIGURE 91- 57 Plasmodium falciparum ring stage in a Giemsa stained blood smear, x 1000.

FIGURE 91- 59 Plasmodium falciparum male and female gametocytes, in a Giemsa stained blood smear, x 1000.

PLASMODIUM VIVAX MALE AND FEMALE GAMETOCYTES

LD BODIES

Salient features

Salient features

1 Male gametocyte (microgametocyte) is spherical and smaller than the female. Cytoplasm stains light blue or pale blue and nucleus is large (Fig. 91-58). 2 Female gametocyte (macrogametocyte) is spherical and large than the male. Cytoplasm is purple and nucleus is small.

1 It is a Giemsa stained thin blood smear showing a red coloured nucleus and pale blue coloured cytoplasm multiple rings in one red blood cell. 2 Leishmania donovani is the causative agent of visceral leishmaniasis or kala-azar (Fig. 91-60). 3 LD bodies, otherwise known as amastigote stage of L. donovani is found mostly inside the macrophages. Some LD bodies are found outside macrophages. 4 L. donovani shows frequent drug resistance to pentavalent antimonials.

FIGURE 91- 58 Plasmodium vivax female gametocyte in a Giemsa stained blood smear, x 1000.

FIGURE 91- 60 LD bodies in a Giemsa stained blood smear, x 1000.

PLASMODIUM FALCIPARUM MALE AND FEMALE GAMETOCYTES

TOXOPLASMA GONDII

Salient features

Salient features

1 Male gametocyte (microgamete) is sickle shaped, broader and shorter. Cytoplasm stains light blue and nucleus is diffuse (Fig. 91-59). 2 Female gametocyte are typically crescent (banana) shaped with rounded or pointed ends. Cytoplasm stains deep blue and nucleus is compact.

1 The typical active multiplying tachyzoites is an important diagnostic form of Toxoplasma gondii (Fig. 91-61). 2 Tachyzoites appear as 3-7 µm, oval to crescent shaped structures with pointed anterior end and rounded posterior end. 3 Tachyzoites can be stained with periodic acid schiff (PAS), Gomori methenamine silver, hematoxylin and eosin, and Wright-Giemsa stain.

Textbook of Practical Microbiology

4 Tissue cyst is the resting form of T. gondii which appear as 40mm to 50mm spherical structure surrounded by eosinophilic, weakly PAS positive cyst wall. It contains hundred of strongly PAS positive, slow growing trophozoites known as bradyzoites.

291

CYST OF ENTAMOEBA HISTOLYTICA/ DISPAR Salient features 1 Iodine wet mount of stool showing quadrinucleate cyst of Entamoeba histolytica/dispar (Fig. 91-63). 2 Cyst contains 4 nuclei, each nucleus has a central karyosome. 3 The cysts of E. histolytica and E.dispar are morphologically similar. 4 The quadrinucleate cyst is the infective form of the parasite. 5 The infection is transmitted by ingestion of water and food contaminated with cysts. 6 It is the causative agent of amoebic dysentery, amoebic liver abscess, etc.

FIGURE 91- 61 Toxoplasma gondii tachyzoites in Wrisht-Giemsa stained blood smear, x 1000.

WUCHERERIA BANCROFTI MICROFILARIA Salient features 1 It is a Giemsa stained thin blood smear showing the sheathed microfilaria. 2 The body of microfilaria shows few nuclei in the body and more distinct tail is tapering and pointed. 3 No nuclei are present in the tail end. 4 Wuchereria bancrofti is the causative agent of lymphatic filariasis (Fig. 91-62). 5 Anopheles, Culex and Aedes mosquito transmit the disease and are the intermediate host. 6 Man is the definitive host.

FIGURE 91- 62 Wuchereria bancrofti microfilaria in a Giemsa stained blood smear, x 1000.

FIGURE 91- 63 Cyst of Entamoeba histolytica / dispar, x 400.

CYST OF GIARDIA INTESTINALIS Salient features 1 Iodine wet mount of stool showing iodine stained brown oval cyst of Giardia intestinalis (Fig. 91-64). 2 Cyst contains four nuclei, and an axostyle which lie diagonally, forming a dividing line within the cyst wall.The cyst is separated from the cyst wall by a clear space. 3 The cyst is the infective form of the parasite. 4 The infection is transmitted by ingestion of water and food contaminated with cysts. 5 It is the causative agent of diarrhoea, malabsorption , etc.

FIGURE 91- 64 Cyst of Giardia intestinalis, x 400.

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Identification of Common Viva Spots

EGG OF ROUND WORM

EGG OF ENTEROBIUS VERMICULARIS

Salient features

Salient features

1 Bile stained egg of Ascaris lumbricoides in the saline wet mount of stool (Fig. 91-65). 2 Embryonated egg is the infective form of the parasite. 3 A. lumbricoides infection is transmitted by faeco -oral transmission. 4 It is the causative agent of intestinal ascariasis.

1 Saline wet mount of stool showing plano-convex and nonbile stained egg of Enterobius vermicularis (Fig. 91-67). 2 Egg is the infective form of the parasite. 3 E. vermicularis infection is transmitted by faeco-oral transmission. 4 It is the causative agent of pruritus ani in children. 5 Auto infection is characteristic of E. vermicularis infection.

FIGURE 91-67 Egg of Enterobius vermicularis, x 400.

EGG OF HYMENOLEPIS NANA FIGURE 91-65 Egg of round worm, x 100.

EGG OF HOOK WORM Salient features 1 Saline wet mount of stool showing non- bile stained egg of hook worms: Ancylostoma duodenale and Necator americanus (Fig. 91-66). 2 The eggs of A. duodenale and N. americanus are morphologically similar. 3 Filariform larva is the infective form of the parasite. 4 Hook worm infection is transmitted by filariform larva piercing the intact skin. 5 It is the causative agent of microcytic hypochromic anemia and tropical pulmonary eosinophilia, etc.

Salient features 1 Saline wet mount of stool showing non-bile stained egg of Hymenolepis nana (Fig. 91-68). 2 Egg contains oncosphere with three pairs of hooklets. 3 Egg is the infective form of the parasite. 3 H. nana infection is transmitted by faeco -oral transmission. 4 It is the causative agent of hymenolepiasis

FIGURE 91-68 Egg of Hymenolepis nana, x 400.

EGGS OF TRICHURIS TRICHIURA Salient features

FIGURE 91-66 Egg of Hook worm, x 400.

1 Saline wet mount of stool showing bile stained egg of Trichuris trichiura (Fig. 91-69). 2 T. trichiura egg is barrel – shaped with mucous plug at each pole. 3 Egg is the infective form of the parasite.

Textbook of Practical Microbiology

3 T. trichiura infection is transmitted by faeco-oral transmission. 4 It is the causative agent of gastrointestinal infections. Heavy infection may complicate as appendicitis.

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3 They cause opportunistic infection especially in patients with HIV. 4 They cause oral thrush, vaginitis, onychonychia etc., in immunocompetent patients.

GERM TUBE TEST Salient features

FIGURE 91- 69 Egg of Trichuris trichura, x 400.

CANDIDA ALBICANS Salient features

1 The test is also called Reynold – Braude phenomenon (Fig. 91-71). 2 It is used to identify and differentiate C. albicans from other Candida species 3 The test is performed by incubating Candida in patients serum at 37°C for 2hours 4 Germ tube production is due to the formation of pseudohyphae by the fungus.

1 It is a yeast-like fungus. 2 In Gram stained smear, C. albicans appear purple in colour (Fig. 91-70)

FIGURE 91- 70 Candida albicans in a Gram’s stained smear, x 1000.

FIGURE 91- 71 Germ tube test, x 400.

FURTHER READINGS 1 2 3 4

Forbes BA, Sahm DF and Weissfeld AS. Bailey and Scott’s Diagnostic Microbiology. 11 th ed. (The CV Mosby Company, St. Louis) 2002. Mackie and McCartney. Practical Medical Microbiology. 14th Edition. Churchill Livingstone. pp. 921, 1996. Parija SC. Textbook of Medical Parasitology. All India Publishers and Distributors. 3nd Edition. 2006. Parija SC. Stool Microscopy. BPKIHS, Dharan, Nepal, 1998.

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Index

A Abbe condenser 4 ABO system 110 Acetoin 78 Acetylcholinesterase 140 Acid fast staining method 27 Acid-alcohol 27 Acid-fast staining 27 Learning Objectives 27 Introduction 27 Principle 27 Requirements 27 Equipments 27 Reagents and glass wares 27 Preparation of strong carbol fuchsin 27 Preparation of 20% sulphuric acid 27 Preparation of 95% alcohol 27 Preparation of acid-alcohol decolouriser 28 Specimen 28 Procedure 28 Quality Control 28 Observation 28 Results and Interpretation 28 Key Facts 29 Viva 30 Acid-fast Staining of Stool Smears 198 Learning Objectives 198 Introduction 198 Principle 198 Requirements 198 Equipments 198 Reagents and lab wares 198 Specimen 198 Procedure 198 Quality Control 198 Observations 199 Results and Interpretation 199 Key Facts 199 Viva 200 Adult hamster 265 Adult mice 259 Adult rabbit 261, 262 Aedes aegypti 269, 268, 291 Aerotolerant anaerobes 52 Aesculin 173 Agar 274 Agar dilution method 97, 98 Learning Objectives 97

Introduction 97 Principle 97 Requirements 97 Equipments 97 Reagents and lab wares 97 Preparation of stock solutions of antibiotics 97 Specimens 97 Preparation of suspension of bacteria 97 Procedure 97 Test procedure 98 Quality Control 98 Observations 98 Results and Interpretation 98 Key Facts 98 Viva 99 Agarose 130 Agglutination 108, 117 Albert’s stain 31, 32, 33, 175, 176, 288 Albert’s staining 31 Learning Objectives 31 Introduction 31 Principle 31 Requirements 31 Equipments 31 Reagents and glass wares 31 Preparation of Albert’s stain I 31 Preparation of Albert’s stain II 31 Specimen 31 Procedure 31 Quality Control 32 Observation 32 Results and Interpretation 32 Viva 32 Key Facts 33 Alcohol 27, 60 Alkaline peptone water 46, 181 Alkaline phosphatase 138, 140 Allantoic cavity 243 Allantoic sac 244 Allergic respiratory distress 269 Alsever’s solution 121 Amido black 126, 131 Amniotic sac 243, 244 Amoebiasis 269 Amoebic antigens 130 Amoebic dysentery 291 Amoebic liver abscess 291 Anaerobic bacilli 54

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Index

Anaerobic bacteria 277 Anaerobic cocci 54 Ancylostoma duodenale 292 Animals and their uses in the laboratory 263 Learning Objectives 263 Introduction 263 Principle 263 Requirements 263 Equipments 263 Reagents and animals 263 Specimen 263 Procedure 263 Quality Control 264 Observations 264 Results and Interpretation 264 Key Facts 266 Viva 266 Anopheles 268, 289 291 Anthrax 269 Antibiotic resistance 157 Antibiotic sensitivity 92 Antibiotic susceptibility testing 278 Antigen 133 Antimicrobial agent 93, 94, 99, 100 Antiseptic 59 Antiseptics and Disinfectants 59 Learning Objectives 59 Introduction 59 Principle 59 Requirements 59 Equipments 59 Reagents and media 59 Specimen 59 Procedure 59 Quality Control 59 Observation 59 Results and Interpretation 59 Key Facts 60 Viva 60 antiseptics 59, 60 Antiseptics 60 Anti-Streptolysin O (ASLO) Test 121 Learning Objectives 121 Introduction 121 Principle 121 Requirements 121 Equipments 121 Reagents and lab wares 121 Specimen 121 Procedure 121 Serum dilution 121 Test procedure 121 Quality Control 121 Observations 121 Results and Interpretation 122 Key Facts 122 Viva 122 Antony Von Leeuwenhoek 2 Argas persicus 270 Arginine dihydrolase test 185 Armadillo 288 Asbestos filters 56 Ascaris lumbricoides 206, 292 ASLO 122 Aspergillus flavus 235 Aspergillus fumigatus 234 Aspergillus niger 235 Aspergillus species 6

Auramine O 30 Autoclave 285 Autolysis 169, 171 Auxotrophs 144 Axenic culture medium 208, 209

B Babes Ernst granules 31 Babesia 204 Babesia species 203 Bacilli 6 Bacillus anthracis 6, 287 Bacitracin test 172 Bacterial Agglutination Test 108 Learning Objectives 108 Introduction 108 Principle 108 Requirements 108 Reagents and glass wares 108 Specimen 108 Procedure 108 Quality Control 109 Observations 109 Results and Interpretation 109 Positive agglutination 109 Negative agglutination 109 Auto agglutination 109 Key Facts 109 Viva 109 Bacterial agglutination tests 109 Bacterial conjugation 155 Learning Objectives 155 Introduction 155 Hfr strains 155 F’ factors and sexduction 155 Principle 155 Requirements 155 Equipments 155 Reagents and glass wares 155 Specimen 155 Procedure 155 Quality Control 156 Observations 156 Results and Interpretation 156 Key Points to Remember 156 Viva 157 Bacterial endospores 38 Bacterial plasmids 145 Bactericidal drugs 93, 94, 101 Bacteriostatic drug 93, 94, 101 Balantidium coli 6, 249 Bancroftian filariasis 269 Barbitone buffer 126 Bartonella quintana 270 Basal media 45, 46 Basic dye 215 Basic stains 20 Bicarbonate buffer 50 Bijou bottle 283 Bile aesculin test 173 Bile solubility test 169 Biological false positive (BFP) reactions 123, 125 Biotin-avidin ELISA 141 Bismuth sulfite agar 86 Blastomyces dermatitides 6 Blocking solution 139 Blood agar 44, 45, 46, 161, 162, 163, 164, 165, 184, 274 Blood group antigens 110

Textbook of Practical Microbiology Blood group antisera 110 Blood grouping 110 Learning Objectives 110 Introduction 110 Principle 110 Requirements 110 Reagents and lab wares 110 Specimen 110 Procedure 110 Quality Control 110 Observations 110 Results and Interpretation 110 Key Facts 111 Viva 111 Blood parasites 201 Body louse 270 Boeck and Drbohlav’s medium 208 Borrelia duttoni 270 Borrelia recurrentis 270 Bound coagulase 68, 69 Boutonneuse fever 271 Box 1-1 Terminology 5 Box 1-2 Size of Different Organisms 6 Box 12-1 List of most common capsulated organisms that can be demonstrated by negative staining 41 Box 17-1 List of Anaerobic Bacilli and Cocci 54 Box 18-1 Pasteurisation 58 Box 19-1 Commonly used disinfectants and their mechanism of actions 60 Box 2-1 Principle of Dark Ground Microscopy 8 Box 20-1 Uses of catalase Test 63 Box 20-2 Catalase Test for Mycobacteria 63 Box 22-1 Free Coagulase 69 Box 29-1 List of media used for detecting production of hydrogen sulphide 86 Box 31-1 Glossary of terms 93 Box 4-1 Demonstrating Motility of Anaerobic Bacteria 12 Box 43-1 VDRL-ELISA 125 Box 43-2 Biological False Positive Reactions of VDRL Test 125 Box 47-1 Reverse Passive Haemagglutination test 133 Box 49-1 1st, 2nd and 3rd Generation ELISA 140 Box 49-2 Dot ELISA 140 Box 5-1 Terminology 17 Box 50-1 Role of Plasmids in drug resistance 146 Box 51-1 Advantages and Disadvantages of Page 150 Box 52-1 Point Mutations and Large Scale Mutations 153 Box 52-2 Mechanisms of Mutation 154 Box 52-3 The Importance of Mutation 154 Box 53-1 Mechanisms of DNA Transfer 156 Box 54-1 Beneficial Effects of Normal Flora 161 Box 6-1 Simple stains and their uses in microbiology laboratory 21 Box 61-1 Identification of Escherichia Coli 179 Box 61-2 Identification of Klebsiella species 179 Box 62-1 Identification of Vibrio Cholerae 182 Box 63-1 Identification of Pseudomonasaeruginosa 185 Box 67-1 Acid Fast Parasites and Parasitic Components 199 Box 68-1 The Parasites found in the Peripheral Blood Smear 203 Box 69-1 Advantages and Disadvantages of the Concentration Methods 207 Box 71-1 Sabouraud’s Dextrose Agar 214 Box 77-1 Predisposing Factors for Candidiasis 226 Box 8-1 Different Modifications of Acid Fast Stain and their uses 29 Box 8-2 Frequently examined specimens for the Detection of Mycobacterium Tuberculosis 29 Box 87-1 Uses of Mice in Laboratory 259 Box 88-1 Use of Rabbits in Laboratory 262 Box 89-1 Use of Laboratory Animals 264 Box 9-1 Rapid staining by direct fluorescent antibody method 32 Box 90-1 Diseases Transmitted by Sand Fly 271

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Box 90-2 Diseases Transmitted by Hard Ticks 271 Box 7-1 Modifications of Gram’s staining 25 Box 7-2 Uses of Gram’s Staining 25 Brain heart infusion agar 46 Bright field microscopy 2 Brilliant green bile broth 249 Bromothymol blue 81, 281 Broth dilution agar method 101 Broth dilution method 100 Learning Objectives 100 Introduction 100 Principle 100 Requirements 100 Equipments 100 Reagents and lab wares 100 Preparation of stock solutions of antibiotics 100 Specimens 100 Preparation of suspension of bacteria 100 Procedure 100 Quality Control 100 Observations 101 Results and Interpretation 101 Key Facts 101 Viva 101 Browne’s tube 56, 57 Brownian movement 12 Brucellosis 253 Buffered distilled water 201 Buffered methylene blue 191

C CAMP (Christie, Atkins and Munch-Peterson) test 173 Candida albicans 6, 215, 293 Candida albicans on Sabourauds dextrose agar 280 Candida species 225 Candle jar 51 Candling 244 Capillary tube method 11, 12 Capsular antigens 36 Capsulated bacteria 36 Capsule 36, 286 Capsule Staining 3 4 Learning Objectives 34 Introduction 34 Principle 34 Requirements 34 Equipments 34 Reagents and glass wares 34 Specimen 34 Procedure 34 For positive staining of smears 34 For negative staining of smears 35 Quality Control 35 Observation 35 Observation of positive staining method 35 Observation of negative staining method 35 Results and Interpretation 35 Positive staining method 35 Negative staining method 35 Key Facts 35 Viva 36 Carbohydrate assimilation 229 Carbohydrate Assimilation Test 229, 230 Learning Objectives 229 Introduction 229 Principle 229 Requirements 229 Equipments 229

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Index

Reagents and lab wares 229 Specimens 229 Procedure 229 Quality Control 229 Observation 229 Results and Interpretation 229 Key Facts 230 Viva 230 Carbohydrate Fermentation Test 231, 232, 280 Learning Objectives 231 Introduction 231 Principle 231 Requirements 231 Equipments 231 Reagents 231 Preparation of indicator broth medium 231 Specimen 231 Procedure 231 Quality Control 231 Observations 232 Results and Interpretation 232 Key Facts 232 Viva 232 Carbohydrate fermentation tests 231 Carbol fuchsin 27, 198 Carbonate buffer 139 Carbonic acid 71 Cardiolipin antigen 123, 125 Cassette ELISA 141 Catalase 62, 64 Catalase negative bacteria 63 Catalase positive bacteria 63 Catalase test 62, 185 Learning Objectives 62 Introduction 62 Principle 62 Requirements 62 Reagents and glass wares 62 Specimen 62 Procedure 62 Slide method 62 Tube method 62 Quality Control 63 Positive control 63 Negative control 63 Observations 63 Slide method 63 Tube method 63 Results and Interpretation 63 Key Facts 64 Viva 64 Catalyst 53 Cell culture 240, 242 Cell lines 240 Cephalosporium 236 Cerebral malaria 289 Cetrimide agar 46, 185 Chick cell agglutination 181 Chick Martin test 60 Chick RBCs agglutination 182 Chikungunya fever 269 Chlorhexidine 60 Chlorination 248 Chocolate agar 44, 161, 163, 275 Cholera 181, 269 Chorioallantoic membrane (CAM) 243, 244 Christensen’s urea agar 71, 227, 281 Chromoblastomycosis 6

CIEP test 130, 131 Citrate negative bacteria 81 Citrate positive bacteria 81 Citrate sulfide agar 86 Citrate utilisation test 80, 81 Learning Objectives 80 Introduction 80 Principle 80 Requirements 80 Equipments 80 Reagents and lab wares 80 Specimen 80 Procedure 80 Quality Control 81 Positive control 81 Negative control 81 Observations 81 Results and Interpretation 81 Key Facts 81 Viva 81 Citrate utilization test 185, 281, 282 Cladosporium 236 CLED medium 46, 178, 180 Clostridium perfringens 248, 287 Co-agglutination test 114, 115 Learning Objectives 114 Introduction 114 Principle 114 Requirements 114 Reagents and lab wares 114 Specimen 114 Procedure 114 Quality Control 114 Observations 114 Results and Interpretation 114 Key Facts 115 Viva 115 CO 2 incubator 160 Coagulase 68 Coagulase negative bacteria 69 Coagulase negative staphylococci 168 Coagulase positive bacteria 69 Coagulase reacting factor (CRF) 69 Coagulase test 68, 166, 167 Learning Objectives 68 Introduction 68 Principle 68 Bound coagulase 68 Free coagulase 68 Requirements 68 Reagents and lab wares 68 Specimen 68 Procedure 68 Slide test 68 Tube test 68 Quality Control 69 Positive control 69 Negative control 69 Observation 69 Results and Interpretation 69 Key Facts 69 Viva 70 Coccidian parasites 198 Coccidioides immitis 6 Cold agglutination test 120 Coliforms 248, 251 Collection of Blood from the Marginal Ear Vein of Rabbit 261 Learning Objectives 261

Textbook of Practical Microbiology Introduction 261 Principle 261 Requirements 261 Laboratory wares 261 Reagents 261 Specimen 261 Procedure 261 Quality Control 261 Observations 261 Results and Interpretation 261 Key Facts 262 Viva 262 Colorado tick fever 271 Competitive ELISA 138, 141 Compound Microscope 3 Learning Objectives 3 Introduction 3 Parts of the compound microscope 3 Microscope stand 3 Main tube 3 Body and arm 3 Substage 3 Foot 3 Stage 3 Microscope optics 4 Mechanical adjustment of a microscope 4 Coarse and fine focusing adjustments 4 Condenser adjustment 4 The light source 4 Principle 4 Magnification 4 Principle involved in the magnification of the object 4 Importance of numerical aperture 5 Requirements 5 Equipment 5 Reagents 5 Specimen 5 Procedure 5 Observations 5 Results and Interpretation 5 Key Facts 6 Viva 6 Compound light microscope 9 Concentration of stool 205, 207 Concentration of Stool for Parasites 205 Learning Objectives 205 Introduction 205 Principle 205 Requirements 205 Equipments 205 Reagents and lab wares 205 Specimen 205 Procedure 206 Saturated salt solution flotation method 206 Formalin-ether sedimentation method 206 Quality Control 206 Observations 206 Results and Interpretation 206 Key Facts 207 Viva 207 Condenser 4 Conjugation 155, 157 Conjunctivitis 269 Coomassie staining 149 Corynebacterium diphtheriae 31, 175, 275 Corynebacterium diphtheriae on potassium tellurite agar 279 Counter-current immunoelectrophoresis (CIEP) 130, 149 Counter-current Immunoelectrophoresis Test 130

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Learning Objectives 130 Introduction 130 Principle 130 Requirements 130 Equipment 130 Reagents and glass wares 130 Specimen 130 Preparation of Veronal buffer 0.075 M (pH 8.6) 130 Preparation of agarose 130 Procedure 130 Quality Control 131 Observations 131 Results and Interpretation 131 Key Facts 131 Viva 131 Coxsackie A and B viruses 258 Craigie’s tube method 11 Crimean - congo hemorrhagic fever 271 Cryptococcal antigen 130 Cryptococcus neoformans 6, 221 Cryptosporidium parvum 198, 199, 200, 258 Culex 269, 291 Cultivation of Fungi 213 Learning Objectives 213 Introduction 213 Principle 213 Requirements 213 Equipments 213 Reagents and lab wares 213 Specimen 213 Procedure 213 Quality Control 213 Observations 213 Results and Interpretation 213 Key Facts 214 Viva 214 Cultivation of Viruses in the Cell lines 240 Learning Objectives 240 Introduction 240 Principle 240 Requirements 240 Equipments 240 Reagents and lab wares 240 Specimen 240 Procedure 240 Quality Control 241 Observations 241 Results and Interpretation 241 Key Facts 242 Viva 242 Cultivation of Viruses in Embryonated Egg 243 Learning Objectives 243 Introduction 243 Principle 243 Requirements 243 Equipments 243 Reagents and lab wares 243 Specimen 243 Procedure 243 Quality Control 244 Observations 244 Results and Interpretation 244 Key Facts 244 Viva 244 Culture media 274 Culture of anaerobic bacteria 53 Learning Objectives 53 Introduction 53

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Index

Principle 53 Requirements 53 Equipments 53 Reagents and media 53 Specimen 53 Procedure 53 Quality Control 54 Observations 54 Results and Interpretation 54 Viva 54 Key Facts 55 Culture of Stool for Entamoeba histolytica 208 Learning Objectives 208 Introduction 208 Principle 208 Requirements 208 Equipments 208 Reagents and lab wares 208 Specimen 208 Procedure 209 Quality Control 209 Observations 209 Results and Interpretation 209 Key Facts 209 Viva 209 Cutaneous mycoses 212 Cyclops 270 Cyclospora cayetanensis 195, 198, 199, 200 Cyst of Entamoeba coli 190193, 196, 206 Cyst of Entamoeba histolytica/dispar 190, 193, 196, 206, 291 Cyst of Giardia intestinalis 190, 193, 196, 206, 291 Cytopathic effect (CPE) 240, 241

D D’ Antonie’s iodine 192 Dark ground microscopy 2, 7, 11 Learning Objectives 7 Introduction 7 Principle 7 Requirements 7 Equipments 7 Reagents 7 Specimen 7 Procedure 7 Observations 7 Results and Interpretation 7 Key Facts 8 Viva 8 Decolourising agent 26, 27, 215 Dengue haemorrhagic fever 268 Deoxycholate citrate agar (DCA) medium 44, 86, 181 Dettol 60 Diarrhea 269, 291 Differential coliform test 249, 250 Differential media 46 Differential stain 21, 26 Diffusion tests 97 Dilute carbol fuchsin 215 Diphtheroids 163 Diphyllobothriasis 271 Diphyllobothrium latum 271 Direct fluorescent antibody test 32, 135, 136 Direct plate technique 65 Disc diffusion test 94 Disinfectants 59, 60 Disinfection 59, 60 DNAase test 167 Dobell and O’Connor’s iodine 192

Dobell’s iodine 192 Dot ELISA 139, 140 Double diffusion method 126 Dracunculiasis 271 Dracunculus medinesis 6 Draughtsman’s colonies 169 Drbohlav’s Locke-egg-serum (LES) medium 208 Dry heat sterilization 285 Durham’s tube 231, 250 Dysentery 269

E E-test 103 Earthenware candles 56 Echinococcus granulosus 283 EDTA 69 Egg of Ascaris lumbricoides 190, 193, 196 Egg of Enterobius vermicularis 292 Egg of hook worm 206, 292 Egg of Hymenolepis nana 292 Egg of round worm 292 Egg of Trichuris trichiura 190, 193, 196, 292 Eggs of Taenia saginata 199 Ehrlich’s reagent 74, 75 Eijkman test 249, 251 Electro-immuno transfer blot (EITB) 150 Electroimmunodiffusion 127, 149 Electron microscopes 2 Electrophoresis 148 Embryonated eggs 243 Endospores 37 Enriched media 45, 46, 275 Enrichment media 46 Entamoeba histolytica 8, 208, 248 Enteric fever 116, 118 Enterobius vermicularis 292 Enzyme-linked immunosorbent assay 138, 141 Learning Objectives 138 Introduction 138 Principle 138 The sandwich ELISA for antigen 138 The indirect ELISA for antibodies 138 Competitive ELISA for antibodies 138 Indirect ELISA to Detect Antibodies 139 Requirements 139 Equipments 139 Reagents and glass wares 139 Carbonate buffer (pH 9.6) 139 Washing buffer (PBS-Tween 20) 139 Conjugate 139 Citric acid phosphate buffer (0.1M) pH 5.0 139 Substrate 139 Specimen 139 Procedure 139 Quality Control 139 Observations 139 Results and Interpretation 139 Sandwich ELISA to Detect Antigen 139 Requirements 139 Equipments 139 Reagents and glass wares 139 Specimen 139 Procedure 139 Quality Control 140 Observations 140 Results and Interpretation 140 Key Facts 141 Viva 141

Textbook of Practical Microbiology Enzymes 138 Epidemic relapsing fever 270 Epidemic typhus 270 Epidermophyton floccosum 236 Epsilometer test (E-test) 102 Learning Objectives 102 Introduction 102 Principle 102 Requirements 102 Reagents and lab wares 102 Specimens 102 Procedure 102 Opening an E-test package 102 Application of strips 102 Quality Control 103 Observations 103 Results and Interpretation 103 Key Facts 103 Viva 103 Escherichia coli 178 Escherichia coli on MacConkey agar 279 Ether 205 Ethidium bromide solution 145 Eye pieces 4

F F factor 155 Facultative anaerobes 52, 54 Faecal Escherichia coli 248 Faecal Streptococci 248 Falciparum malaria 289 Filariform larva 292 Floatation method 207 Fluorescence microscopy 2 Fluorescent microscope 135, 136 Fluorescent staining 30 Foetal calf serum (FCS) 240 Forage mite 269 Formalin 205 Formalin-ether sedimentation method 205, 206, 207 Free coagulase 68 Fungal elements 219 Fusarium 234

G Gas gangrene 38 Gaspak system 53 Gel diffusion test 127 Gelatin hydrolysis test 176 Germ tube test 225, 226, 293 Learning Objectives 225 Introduction 225 Principle 225 Requirements 225 Equipments 225 Reagents and glass wares 225 Specimens 225 Procedure 225 Quality Control 225 Observations 225 Results and Interpretation 226 Key Facts 226 Viva 226 Germ tubes 225, 226 Giardia species 248 Giemsa stains 201, 203 Glucose 6-phosphate dehydrogenase 140 Glucose broth 277

Glucose phosphate medium 82 Glucose with Durham’s tube 281 Glutaraldehyde 60 Glycerol 195, 218 Gnathostoma hispidium 271 Gnathostoma spinigerum 271 Goat antimouse immunoglobulin 139 Gomori methenamine silver 290 Gram negative cocci 24 Gram positive bacilli 24 Gram positive bacteria 24 Gram positive cell wall 26 Gram positive cocci 24, 286 Gram variable bacteria 26 Gram’s iodine 215 Gram’s iodine 23 Gram’s stain 26, 37, 161, 162, 164, 215 Gram’s Staining for Fungi 215 Learning Objectives 215 Introduction 215 Principle 215 Requirements 215 Equipments 215 Reagents and lab wares 215 Specimen 215 Procedure 215 Preparation of fungal smear 215 Staining Procedure 215 Quality Control 215 Observation 216 Results and Interpretation 216 Key Facts 216 Grams staining 23 Learning Objectives 23 Introduction 23 Principle 23 Requirements 23 Equipments 23 Reagents and glass wares 23 Preparation of methyl violet stain 23 Preparation of Gram’s iodine 23 Preparation of 1% safranine 23 Specimen 24 Preparation of bacterial smear 24 Preparation of bacterial smear 24 Procedure 24 Quality Control 24 Observation 24 Results and Interpretation 24 Key Facts 26 Viva 26 Group A streptococci 172 Group B streptococci 172 Group D streptococci 172 Guinea pig 263, 264, 265

H H2S producing bacteria 86 Haemoparasites 201 Haemophilus influenzae 287 Hamsters 263, 265 Hanging drop preparation 11, 181 Learning Objectives 11 Introduction 11 Principle 11 Requirements 11 Equipments 11 Reagents and glass wares 11

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Index

Specimen 11 Procedure 11 Quality Control 11 Observations 12 Results and Interpretation 12 Key Facts 13 Viva 13 Hard ticks 270 Head louse 270 Heat stable catalase test 63 Hektoen enteric agar 86 HeLa 240 Hemolysis on blood agar 167 Hep 2 240 Hepatitis B antigen 130 Heterophile agglutination tests 107 Histoplasma capsulatum 6 Holder method 58 Hook worm egg 190, 193, 196 Horseradish peroxidase 138, 140 Hot air oven 58, 285 House dust mite 269 House flies 269 Hungate procedure of anaerobiosis 54 Hybridization probes 144 Hydatid antigens 130, 132 Hydatid cyst 283 Hydatid fluid 283 Hydrogen Sulfide Test 85 Learning Objectives 85 Introduction 85 Principle 85 Requirements 85 Equipments 85 Reagents and lab wares 85 Specimen 85 Procedure 85 Quality Control 86 Positive control 86 Negative control 86 Observations 86 Results and Interpretation 86 Viva 86 Key Facts 87 Hymenolepis nana 6, 292

I Identification of Common Fungi 233 Learning Objectives 233 Introduction 233 Principle 233 Requirements 233 Equipments 233 Reagents and lab wares 233 Specimen 233 Procedure 233 Quality Control 233 Observation 233 Results and Interpretation 233 Rhizopus 233 Colony morphology 233 Microscopy 234 Mucor 234 Colony morphology 234 Microscopy 234 Alternaria 234 Colony morphology 234 Microscopy 234

Fusarium 234 Colony morphology 234 Microscopy 234 Aspergillus fumigatus 234 Aspergillus niger 235 Aspergillus flavus 235 Penicillium species 235 Conidiophores 235 Phialids 235 Cladosporium 236 Colony 236 Microscopy 236 Cephalosporium 236 Colony 236 Microscopy 236 Trichophyton verrucosum 236 Colony 236 Trichophyton violaceum 236 Colony 236 Epidermophyton floccosum 236 Colony 236 Viva 237 Identification of Common Insects 268 Learning Objectives 268 Introduction 268 Principle 268 Requirements 268 Equipments and lab wares 268 Specimen 268 Mosquitoes 268 General features 268 Identifying features of Anopheles 268 Diseases transmitted 268 Identifying features of Aedes 268 Diseases transmitted 268 Identifying features of Culex 269 Diseases transmitted 269 Sand Fly 269 General features 269 House Flies 269 Identifying features 269 Diseases transmitted 269 Itch Mite 269 Identifying features 269 Diseases transmitted 269 Trombiculid Mite 269 Identifying features 269 Diseases transmitted 269 Hard Tick 270 Identifying features 270 Soft Tick 270 Identifying features 270 Diseases transmitted 270 Rat Flea 270 Identifying features 270 Louse 270 Identifying features 270 Diseases transmitted 270 Cyclops 270 Identifying features 270 Viva 271 Identification of Common Viva Spots 274 Learning Objectives 274 Culture Media 274 Nutrient Agar 274 Composition 274 Preparation 274 Modifications 274

Textbook of Practical Microbiology Uses 274, 275, 276, 277, 278, 283, 284, 285 Blood Agar 274 Composition 274 Preparation 275 Chocolate Agar 275 Composition 275 Preparation 275 MacConkey Agar 275 Composition 275 Preparation 275 Loeffler’s Serum Slope 275 Composition 276 Preparation 276 Lowenstein-Jensen (LJ) Medium 276 Composition 276 Preparation 276 Robertson Cooked Meat (RCM) Broth 276 Composition 276 Preparation 276 Sabouraud’s Dextrose Agar (SDA) 277 Composition 277 Preparation 277 Peptone water 277 Composition 277 Preparation 277 Glucose Broth 277 Composition 278 Preparation 278 Culture Media with Growth 278 Streptococcus pyogenes on Blood Agar 278 Salient features 278 Staphylococcus aureus on Nutrient Agar 278 Salient features 278 Proteus spp. on Blood Agar 278 Salient features 278 Pseudomonas aeruginosa on Nutrient Agar 279 Salient features 279 Escherichia coli on MacConkey Agar 279 Salient features 279 Klebsiella spp. on MacConkey Agar 279 Salient features 279 Corynebacterium diphtheriae on Potassium Tellurite Agar 279 Salient features 279 Mycobacterium tuberculosis on Lowenstein Jensen (LJ) Medium 280 Salient features 280 Candida albicans on Sabouraud’s Dextrose Agar (SDA) 280 Salient features 280 Biochemical Reactions 280 Carbohydrate Fermentation Tests 280 Salient features 280 Glucose with Durham’s Tube 281 Salient features 281 Indole Test 281 Salient features 281 Urease Test 281 Salient features 281 Citrate Utilization Test 281 Salient features 281 Phenyl Pyruvic Acid Test (PPA) 282 Salient features 282 Triple Sugar Iron (TSI) Agar 282 Salient features 282 Specimens of Parasite 282 Ascaris lumbricoides Adult Worm 282 Salient features 282 Hydatid Cyst 283 Salient features 283 Glass Wares 283

Universal Container 283 Bijou Bottle 283 Tuberculin Syringe 283 Graduated Pipette 284 Pasteur Pipette 284 NIH Swab 285 Incubator 285 Salient features 285 Hot Air Oven 285 Salient features 285 Autoclave 285 Salient features 285 Microscopy Slides 286 Gram Positive Cocci 286 Salient features 286 Streptococcus pneumoniae 286 Salient features 286 Neisseria gonorrhoeae 286 Salient features 286 Gram Negative Bacilli 286 Salient features 286 Haemophilus influenzae 287 Salient features 287 Vibrio cholerae 287 Salient features 287 Bacillus anthracis 287 Salient features 287 Clostridium perfringens 287 Salient features 287 Treponema pallidum 287 Salient features 287 Albert staining 288 Salient features 288 Ziehl-Neelsen Staining for Mycobacterium tuberculosis 288 Salient features 288 Ziehl-Neelsen Staining for Mycobacterium leprae 288 Salient features 288 Negri Bodies 288 Salient features 288 Multinucleate Giant Cells Measles 289 Salient features 289 Molluscum Bodies 289 Salient features 289 Plasmodium vivax Ring Stage 289 Salient features 289 Plasmodium falciparum Ring Stage 289 Salient features 289 Plasmodium vivax Male and Female Gametocytes 290 Salient features 290 Plasmodium falciparum Male and Female Gametocytes 290 Salient features 290 LD Bodies 290 Salient features 290 Toxoplasma gondii 290 Salient features 290 Wuchereria bancrofti Microfilaria 291 Salient features 291 Cyst of Entamoeba histolytica/dispar 291 Salient features 291 Cyst of Giardia intestinalis 291 Salient features 291 Egg of Round Worm 292 Salient features 292 Egg of Hook Worm 292 Salient features 292 Egg of Enterobius vermicularis 292 Salient features 292 Egg of Hymenolepis nana 292

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304

Index

Salient features 292 Eggs of Trichuris trichiura 292 Salient features 292 Candida albicans 293 Salient features 293 Germ Tube Test 293 Salient features 293 Identification of Corynebacterium diphtheriae 175 Learning Objectives 175 Introduction 175 Specimen 175 Tests for the identification of Corynebacterium diphtheriae 175 Direct examination 175 Albert’s stain 175 Culture 175 Biochemical tests 175 Key Facts 176 Viva 176 Identification of Lactose Fermenting Enterobacteriaceae 178 Learning Objectives 178 Introduction 178 Specimens 178 Tests for Identification of E. Coli and Klebsiella spp. 178 Direct examination 178 Gram’s stain 178 Culture 178 Biochemical tests 178 Antibiotics susceptibility testing 178 Key Facts 180 Viva 180 Identification of Pseudomonas aeruginosa 184 Learning Objectives 184 Introduction 184 Specimens 184 Tests for Identification of Pseudomonas Aeruginosa 184 Direct examination 184 Culture 184 Oxidase test 184 Biochemical tests 184 Antibiotic susceptibility testing 184 Key Facts 185 Viva 185 Identification of Staphylococcus aureus 166 Learning Objectives 166 Introduction 166 Specimen 166 Tests for the Identification of Staphylococcus aureus 166 Direct examination 166 Culture 166 Coagulase test 166 Deoxyribonuclease test 166 Mannitol salt agar 166 Novobiocin sensitivity 167 Key Facts 168 Viva 168 Identification of Streptococcus pneumoniae 169 Learning Objectives 169 Introduction 169 Specimen 169 Tests for the Identification of Streptococcus pneumoniae 169 Direct examination 169 Culture 169 Bile solubility test 169 Optochin test 169 Inulin fermentation 170 Key Facts 170 Viva 171 Identification of Vibrio cholerae 181

Learning Objectives 181 Introduction 181 Specimens 181 Tests for Identification of Vibrio Cholerae 181 Direct examination 181 Hanging drop preparation 181 Culture 181 Biochemical tests 181 Antibiotic susceptibility testing 181 Key Facts 182 Viva 183 Identification of b-haemolytic Streptococci 172 Learning Objectives 172 Introduction 172 Specimen 172 Tests for Identification of Streptococcus pyogenes 172 Direct examination 172 Culture 172 Bacitracin sensitivity test 172 CAMP (Christie, Atkins and Munch-Peterson) test 173 Bile aesculin test 173 Viva 173 Key Facts 174 IHA test 133, 134, 284 Illuminating source 4 Illumination 5 Immunodiffusion 127 Immunoelectrophoresis test 107, 128, 135 Learning Objectives 128 Introduction 128 Principle 128 Requirements 128 Equipments 128 Reagents and glass wares 128 Specimen 128 Procedure 128 Quality Control 129 Observations 129 Results and Interpretation 129 Key Facts 129 Viva 129 Immunoelectrophoresis 128, 129 Immunofluorescence Test 135 Learning Objectives 135 Introduction 135 Principle 135 Requirements 135 Equipments 135 Reagents and lab wares 135 Specimen 135 Procedure 135 Quality Control 136 Observations 136 Results and Interpretation 136 Key Facts 136 Viva 136 In-use test 60 Incineration 58 Incomplete antibodies 108 Incubators 285 India ink 34, 41, 221, 222, 286 India ink preparation 35, 212, 221, 222 Learning Objectives 221 Introduction 221 Principle 221 Requirements 221 Equipments 221 Reagents and lab wares 221

Textbook of Practical Microbiology Specimen 221 Procedure 221 Quality Control 221 Observations 221 Results and Interpretation 221 Key Facts 222 Viva 222 India-ink method 40 Indicator broth medium (IBM) 231 Indicator medium 275 Indirect ELISA 141 Indirect haemagglutination test 132 Learning Objectives 132 Introduction 132 Principle 132 Requirements 132 Lab wares 132 Reagents 132 Specimen 133 Procedure 133 Sensitisation of chick RBCs with OSD of the antigen 133 Performance of the IHA test 133 Quality Control 133 Observations 133 Results and Interpretation 133 Key Facts 133 Viva 134 Indirect paper strip procedure 65 Indole negative bacteria 74, 75 Indole positive bacteria 74, 75 Indole test 74, 185, 281 Learning Objectives 74 Introduction 74 Principle 74 Requirements 74 Equipments 74 Reagents and lab wares 74 Specimen 74 Procedure 74 Quality Control 74 Positive control 74 Negative control 74 Observation 74 Results and Interpretation 74 Key Facts 75 Viva 75 Infant hamsters 265 Infant rabbit 262 Insects 268 Inspissation 58 Intestinal coccidian parasites 199, 200 Intravenous Inoculation into Mice Tail Vein 258 Learning Objectives 258 Introduction 258 Principle 258 Requirements 258 Equipments 258 Reagents and glass wares 258 Specimen 258 Procedure 258 Procedure for loading the syringe for injection 258 Animal preparation for injection 258 Injection of material 258 Quality Control 259 Observations 259 Results and Interpretation 259 Key Facts 260 Viva 260

305

Inulin fermentation 170 Iodine wet mount 191, 192, 194 Iodine Wet Mount of Stool 192 Learning Objectives 192 Introduction 192 Principle 192 Requirements 192 Equipments 192 Reagents and glass wares 192 Specimen 192 Procedure 192 Quality Control 192 Observations 193 Results and Interpretation 193 Key Facts 194 Viva 194 Iodine wet mount preparation 189, 192 Isoantibodies 111 Isolation of Antibiotic Resistant Mutant 152 Learning Objectives 152 Introduction 152 The importance of mutation 152 Principle 152 Requirements 152 Equipments 152 Reagents and lab wares 152 Specimen 152 Procedure 152 Quality Control 153 Observations 153 Results and Interpretation 153 Key Facts 154 Viva 154 Isolation of Plasmids 145 Learning Objectives 145 Introduction 145 Principle 145 Requirements 145 Equipments 145 Reagents and lab wares 145 Preparation of ethidium bromide stock solution 145 Preparation of ethidium bromide working solution 145 Preparation of Luria-Bertani medium (LB medium) 145 Specimen 145 Procedure 145 Extraction of plasmid 145 Electrophoresis on agarose gel 146 Quality Control 146 Observations 146 Results and Interpretation 146 Key Facts 146 Viva 146 Isolation of Pure Cultures 14 Learning Objectives 14 Introduction 14 Principle 14 Requirements 14 Equipment and labwares 14 Reagents 14 Specimen 15 Procedure 15 For streak plate method 15, 16 For spread plate method 15, 16 For pour plate method 15, 16 Quality Control 15 Observations 15 Results and Interpretation 16 Viva 16

306

Index

Key Facts 17 Isospora belli 198, 199 Itch mite 269

J Japanese encephalitis 269 Jaswant Singh Bhattacharjee stain 203 Jensen’s Gram method for smears 25

K K antigen 36 Köhler illumination 3, 6 Kala azar 271 Kinyoun’s modification of acid-fast stain 29 Kirby-Bauer disc diffusion method 92 Kirby-Bauer method 92, 96 Learning Objectives 92 Introduction 92 Principle 92 Requirements 92 Equipments 92 Reagents and lab wares 92 Preparation of 0.5 McFarland standard 92 Specimens 92 Preparation of suspension of bacteria 92 Procedure 92 Quality Control 93 Observations 93 Results and Interpretation 93 Key Facts 93 Viva 94 Kirby-Bauer’s chart 93 Klebsiella species 178 Klebsiella species on MacConkey agar 279 Kligler’s iron agar median 85, 86, 185 Koch’s postulates 288 KOH wet mount preparation 219, 220 Kopeloff and Beerman’s Gram method 25 Koser’s citrate 80, 81 Koser’s liquid citrate medium 281 Kovac’s reagent 74, 75, 281 Krebs cycle 80 Kyasanur forest disease 271

L Löwenstein-Jensen medium 44 Laboratory animals 263 Lactophenol cotton blue (LPCB) stain 195, 217 Lactophenol Cotton Blue (LPCB) Wet Mount of Fungi 217 Learning Objectives 217 Introduction 217 Principle 217 Requirements 217 Equipments 217 Reagents and lab wares 217 Specimen 217 Procedure 217 Scotch tape preparation 217 Tease mount preparation 217 Quality Control 218 Observations 218 Results and Interpretation 218 Key Facts 218 Viva 218 Lactophenol cotton blue (LPCB) wet mount 212, 217 Lactose fermenting Enterobacteriaceae 178 Lane’s saturated salt solution floatation method 205 Latex agglutination test 112, 113

Learning Objectives 112 Introduction 112 Principle 112 Requirements 112 Reagents and lab wares 112 Specimen 112 Procedure 112 Quality Control 112 Observations 112 Results and Interpretation 113 Key Facts 113 Viva 113 LD bodies 290 Lead acetate agar 85, 86 Leishman stains 201, 203 Leishman’s 201 Leishman’s stain 201 Leishman’s Staining of Peripheral Blood Smears 201 Learning Objectives 201 Introduction 201 Principle 201 Requirements 201 Equipments 201 Reagents and lab wares 201 Preparation of EDTA anticoagulated blood 201 Preparation of Leishman’s stain 201 Specimen 202 Procedure 202 Preparation of thin blood smear 202 Preparation of thick blood smear 202 Preparation of combined thick and thin films 202 Leishman’s staining 202 Quality Control 202 Observations 202 Results and Interpretation 203 Key Facts 203 Viva 204 Leishmania donovani 290 Lepromin test 284 Levaditi stain 287 Light microscopy 2 Listeria monocytogenes 261 Locke’s solution 208 Loeffler’s methylene blue 21, 27, 28 Loeffler’s serum slope 175, 275, 276, 280 Louse 270 LPCB Wet Mount of Stool 195 Learning Objectives 195 Introduction 195 Principle 195 Requirements 195 Equipments 195 Reagents and glass wares 195 Specimen 195 Procedure 195 Quality Control 196 Observations 196 Results and Interpretation 196 Key Facts 197 Viva 197 LPCB wet mount preparation 196 Lugol’s’ iodine 192 Luria-Bertani medium 145 Lyme disease 271 Lymphatic filariasis 291 Lysine decarboxylation test 185 Lysine iron agar 86 Lysol 60

Textbook of Practical Microbiology

M MacConkey agar 45, 46, 178, 184, 275 MacConkey broth 249 Macleod’s potassium tellurite agar media 175 Macrocapsule 34 Macrogametocyte 290 Magnesium sulphate floatation method 206 Magnification 6 Malabsorption 291 Malachite green solution 276 Malachite green stain 37 Malaria 268 Male gametocyte 290 Mannitol salt agar 166, 167 Mc Fadyean reaction 21 McFadyean reaction 35 McFarland standard 92, 95 McIntosh and Fildes jar 53 Measurement of Microorganisms 9 Learning Objectives 9 Introduction 9 Principle 9 Requirements 9 Equipments 9 Reagents 9 Specimen 9 Procedure 9 Quality Control 10 Observations 10 Results and Interpretation 10 Key Facts 10 Viva 10 Media for Routine Cultivation of Bacteria 44 Learning Objectives 44 Introduction 44 Principle 44 Basal media 44 Enriched medium 44 Enrichment media 44 Selective media 44 Differential media 44 Requirements 45 Equipments 45 Reagents and media 45 Specimen 45 Procedure 45 Quality Control 45 Observations 45 Results and Interpretation 46 Key Facts 46 Viva 46 Membrane filtration method 249 Mesocyclops 271 Metachromatic granules 31, 32, 176, 288 Methyl alcohol 201 Methyl red test 76 Learning Objectives 76 Introduction 76 Principle 76 Requirements 76 Equipments 76 Reagents and lab wares 76 Specimen 76 Procedure 76 Quality Control 77 Positive control 77 Negative control 77 Observation 77

307

Results and Interpretation 77 Key Facts 77 Viva 77 Methyl violet stain 23, 215 Methylene blue (counter stain) 198 Methylene blue reduction test 252 Methylene blue test 252 Mice 263 Microbiology of Air 254 Learning Objectives 254 Introduction 254 Principle 254 Requirements 254 Equipments 254 Reagents and lab wares 254 Procedure 254 Quality Control 254 Observations 254 Results and Interpretation 254 Key Facts 255 Viva 255 Microbiology of Water 248 Learning Objectives 248 Introduction 248 Coliforms 248 Faecal or thermotolerant coliforms 248 Faecal Escherichia coli 248 Faecal streptococci 248 Clostridium perfringens 248 Collection of water samples 249 Principle 249 Plate count 249 Detection of coliform bacteria 249 Presumptive coliform test – Multiple tube technique 249 Differential coliform test 249 Membrane filtration method 249 Detection of faecal streptococci 249 Examination for Cl. perfringens 249 Requirements 249 Equipments 249 Reagents 249 Preparation of MacConkey broth 249 Preparation of double strength medium 249 Preparation of single strength medium 249 Preparation of brilliant green bile broth 250 Specimen 250 Procedure 250 Quality Control 250 Observations 250 Results and Interpretation 250 Differential coliform count 250 Key Facts 251 Viva 251 Microbiology of Milk 252 Learning Objectives 252 Introduction 252 Principle 252 Requirements 252 Equipments 252 Reagents and glass wares 252 Specimen 252 Procedure 252 Quality Control 253 Observations 253 Results and Interpretation 253 Key Facts 253 Viva 253 Microcapsule 34

308

Index

Microfilaria of Brugia malayi 203 Microfilaria of Loa loa 203 Microfilaria of Mansonella perstans 203 Microfilaria of Wuchereria bancrofti 203 Miliary tuberculosis 29 Minimum bactericidal concentration (MBC) 100, 101 Minimum inhibitory concentration (MIC) 97, 98, 100, 101, 102 Missense mutations 153 Modified acid fast staining 199 Modified acid fast staining of faeces 198 Modified Ziehl-Neelsen stain 37 Moist heat sterilization 285 Molluscum bodies 289 Monkeys 263, 265 Monoclonal antibodies 139 Mordant 24, 215 Mosquitoes 268 Motile bacteria 12 Mouse 258, 260, 264 Mouse virulence 170 MR negative bacteria 77 MR positive bacteria 77 MR test 76, 77 Mucor 234 Multinucleate giant cells 289 Mutations 152, 154 Mycobacterium leprae 258, 288 Mycobacterium tuberculosis 28, 276 Mycobacterium tuberculosis on Lowenstein- Jensen’s medium 280

N N-N tetramethyl para-phenylene diamine hydrochloride 65 NCCLS QC ranges 103 NCCLS table 98 Necator americanus 292 Negative staining 40, 41 Learning Objectives 40 Introduction 40 Principle 40 Requirements 40 Equipments 40 Reagents and glass wares 40 Specimen 40 Procedure 40 Quality Control 40 Observations 41 Results and Interpretation 41 Key Facts 41 Viva 41 Negri bodies 288 Neisser’s stain 31, 33, 176 Neisseria gonorrhoea 286 Neutral red 180 Neutralization test 107 Nigrosin staining 34, 40 NIH Swab 285 Nitrate agar slant 88 Nitrate broth 89 Nitrate reduction test 88, 89, 185 Learning Objectives 88 Introduction 88 Principle 88 Requirements 88 Equipments 88 Reagents and lab wares 88 Specimen 88 Procedure 88 Quality Control 88

Positive control 88 Negative control 88 Observation 88 Results and Interpretation 88 Key Facts 89 Viva 89 Nitrates 89 Non agglutinable vibrios 183 Non motile bacteria 12 Non-bile stained eggs 193 Non-fermenters 184 Nonsense mutations 153 Normal flora 160 Normal Microbial Flora of the Mouth 160 Learning Objectives 160 Introduction 160 Principle 160 Requirements 160 Equipments 160 Reagents and lab wares 160 Specimen 160 Procedure 160 Quality Control 160 Observations 160 Results and Interpretation 161 Key Facts 161 Viva 161 Normal Microbial Flora of the Skin 164 Learning Objectives 164 Introduction 164 Principle 164 Requirements 164 Equipments 164 Reagents 164 Specimen 164 Procedure 164 Observations 164 Results and Interpretation 164 Viva 164 Key Facts 165 Normal Microbial Flora of the Throat 162 Learning Objectives 162 Introduction 162 Principle 162 Requirements 162 Equipments 162 Reagents and lab wares 162 Specimen 162 Procedure 162 Observations 162 Results and Interpretations 162 Key Facts 163 Viva 163 Normal flora in the throat 162 Normal flora on the skin 164 Normal microbial flora 163 Northern blotting 149 Norwegian itch of man 269 Nosocomial fungal infections 212 Novobiocin sensitivity 167 Novobiocin-sensitive staphylococci 168 Numerical aperture 5, 6 Nutrient agar 45, 46, 184, 274 Nutrient broth 274

O O/129 reagent 183 Obligate aerobes 52

Textbook of Practical Microbiology Obligate anaerobes 52, 53 Ocular micrometer 9 Onychonychia 293 Optimum sensitizing dose (OSD) of the antigen 132 Optochin (ethyl hydrocupreine hydrochloride) 169 Optochin sensitivity 170 Optochin test 169 Oral thrush 293 Oriental sore 271 Orientia tsutsugamushi 269 Ornithodorus lahorensis 270 Ornithodorus species 270 Ornithodorus tholozani 270 Oroya fever 271 Osmk hemorraghic fever 271 Oxidase negative bacteria 66 Oxidase positive bacteria 66 Oxidase reagent 65 Oxidase test 65, 66, 185 Learning Objectives 65 Introduction 65 Principle 65 Requirements 65 Reagents and glass wares 65 Specimen 65 Procedure 65 Direct plate technique 65 Indirect filter paper strip procedure 65 Quality Control 66 Positive control 66 Negative control 66 Observations 66 Direct plate technique 66 Indirect filter paper strip procedure 66 Results and Interpretation 66 Viva 66 Key Facts 67 Oxygen Requirement for Growth of Bacteria 51 Learning Objectives 51 Introduction 51 Principle 51 Aerobes 51 Microaerophiles 51 Obligate anaerobes 51 Aerotolerant anaerobes 51 Facultative anaerobes 51 Requirements 51 Equipments 51 Reagents and media 51 Specimen 51 Procedure 52 Quality Control 52 Observations 52 Results and Interpretation 52 Key Facts 52 Viva 52

P P-dimethyl amino benzaldehyde 75, 281 P-phenylene diamine dihydrochloride 67 Paracoccidioides brasiliensis 6 Parvo virus 6 Passive agglutination tests 106 Pasteur pipette 284 Pasteurisation 56, 58 Paul-Bunnel test 120 Pediculus capitus 270 Pediculus corporis 270

Penicillium species 235 Peptone water 46, 74, 274, 277 Periodic acid schiff (PAS) 290 Permanent staining of blood smear 201 PH requirement of bacteria 49 pH Requirement for Growth of Bacteria 49 Learning Objectives 49 Introduction 49 Principle 49 Requirements 49 Equipments 49 Reagents 49 Specimen 49 Procedure 49 Quality Control 49 Observations 49 Results and Interpretation 49 Key Facts 50 Viva 50 Phase-contrast microscope 2, 38 Phenol 60, 218 Phenol red 82, 281, 282 Phenyl pyruvic acid test 282 Phosphatase test 252 Phosphate buffer 50 Phosphate buffer saline 132 Phthirus pubis 270 Physiological saline 189 Pikes medium 46 Plasmids 145, 146, 157 Plasmodium 204 Plasmodium falciparum 203 Plasmodium Falciparum male and female Gametocytes 290 Plasmodium Falciparum Ring stage 289 Plasmodium malariae 203 Plasmodium ovale 203 Plasmodium vivax 203 Plasmodium Vivax male and female Gametocytes 290 Plasmodium vivax ring stage 289 Pneumococcal antigen 114 Point mutations 152, 153, 154 Polar bodies 31 Polar flagellum 11 Poliomyelitis 269 Polyacrylamide Gel Electrophoresis 148 Learning Objectives 148 Introduction 148 Principle 148 Requirements 148 Equipments 148 Reagents and lab wares 148 Stock solutions 148 Working solutions 148 Separating gel buffer (4x) 148 Stacking gel buffer (4x) 148 10% Ammonium persulfate (APS) 148 Electrophoresis/Running Buffer (1x) 149 Sample buffer 149 Staining solution: 1000ml. 149 Destaining solution: 1000ml. 149 Specimen 149 Procedure 149 Quality Control 149 Observations 149 Results and Interpretation 149 Viva 150 Key Facts 151 Polychrome methylene blue 20, 21

309

310

Index

Polymetaphosphate 175 Polymyxin-B (50U) sensitivity 181, 182 Polysaccharide capsule 36, 279 Polyxenic culture medium 208, 209 Ponder’s stains 176 Positive staining technique 35 Potable water 248 Potassium hydroxide (KOH) wet mount 212, 219 Potassium Hydroxide Wet Mount of Fungi 219 Learning Objectives 219 Introduction 219 Principle 219 Requirements 219 Equipments 219 Reagents and lab wares 219 Specimen 219 Procedure 219 Quality Control 219 Observations 219 Results and Interpretations 219 Key Facts 220 Viva 220 Potassium nitrate broth (KNO3 ) 88 Potassium tellurite blood agar 275, 279, 288 Pour plate inoculation procedure 14 Pour plate method 14 Pox virus 6 PPA test 282 Precipitation tests 107 Preston and Morrell’s Gram method 25 Presumptive coliform count 249, 250 Protein-A ELISA 141 Proteus mirabilis 279 Proteus OX 19, OX 2 and OX K antigens 119, 120 Proteus species on blood agar 278 Proteus vulgaris 279 Prototrophs 144 Protozoal cysts 195, 205 Protozoal infections 201, 204 Prozone reaction 108 Pseudomonas aeruginosa 184 Pseudomonas aeruginosa on nutrient agar 279 Pubic louse 270 Puch’s stain 31, 33 Pulmonary tuberculosis 29 Pure culture 14, 16, 17 Pyocyanin 184 Pyruvic acid 78

Q Q fever 253 Quellung reaction 170

R Rabbit 261, 263, 264 Rabbit plasma 68, 69 Radial immuno diffusion 126 Radial Immunodiffusion Test 126 Learning Objectives 126 Introduction 126 Principle 126 Requirements 126 Equipments 126 Reagents and glass wares 126 Specimen 126 Procedure 126 Preparation of antibody containing gels 126 Calibration of reference graph 126

Testing unknown serum samples 127 Quality Control 127 Observations 127 Results and Interpretation 127 Key Facts 127 Viva 127 Rat Flea 270 Reaginic antibodies 123 Recombinant DNA technology 144 Relapsing fever 270 Resolution 5, 6 Resolving power 5 Restriction endonucleases 144 Reverse passive haemagglutination test 132, 133 Reynold – Braude phenomenon 293 Rheumatoid factor (RF) 112 Rhinosporidium seeberi 6 Rhizopus 217, 233 Rhodamine 135 Rickettesia orientails 269 Rickettsia prowazaki 270 Rickettsial infection 119 Rideal Walker test 60 Rift valley fever 269 Robertson cooked meat (RCM) 53, 276 Robertson’s cooked meat (RCM) medium 52 Robinson’s medium 208 Rocky mountain Spotted fever 271 Romanowsky’s stains 201, 204 Russian spring summer encephalitis 271

S Sabouraud’s dextrose agar (SDA) 164, 213, 214, 215, 225, 229, 277 Safranine 23 Safranine stain 37 Saline Wet Mount of Stool 189 Learning Objectives 189 Introduction 189 Principle 189 Requirements 189 Equipments 189 Reagents and glass wares 189 Specimen 189 Procedure 189 Quality Control 189 Observations 189 Results and Interpretation 190 Key Facts 191 Viva 191 Saline wet mount preparation of faeces 189 Salmonella-Shigella agar 86 Sand Fly 269 Sand fly fever 271 Sandwich ELISA 141 Sarcocystis hominis 199 Sarcoptes scabies var hominis 269 Saturated salt solution flotation method 205, 206, 207 Scabies 269 Schaeffer Fulton method 37, 38 Scolices of Echinococcus granulosus 199 Scotch tape preparation 217 Scrub typhus mite 269 SDS-PAGE 150 Sedimentation method 207 Selenite F broth 45, 46 Seller’s technique 289 Semi quantitative catalase test 63 Sereny’s test 262

Textbook of Practical Microbiology Settle plate method 254, 255 Sheather’s sucrose floatation method 200, 206 Significant bacteriuria 25 Simmon’s citrate agar 280, 281 Simmon’s citrate medium 80, 81 Simple stain 21 Simple staining 20 Learning Objectives 20 Introduction 20 Principle 20 Requirements 20 Equipments 20 Reagents and glass wares 20 Preparation of Loeffler’s methylene blue stain 20 Preparation of polychrome methylene blue 20 Specimen 20 Procedure 20 Quality Control 20 Observations 20 Results and Interpretation 21 Key Facts 21 Viva 21 Sintered glass filter 56 Slide agglutination tests 106, 108 Slide coagulase test 70 Slide culture 223, 224 Slide Culture for Fungi 223 Learning Objectives 223 Introduction 223 Principle 223 Requirements 223 Equipments 223 Reagents 223 Specimen 223 Procedure 223 Quality Control 224 Observations 224 Results and Interpretation 224 Key Facts 224 Viva 224 Slide culture preparation 217 Slide culture set 223 Slide flocculation test 123 Slide test 68 Slit sampler method 254, 255 Sodium dodecyl sulfate - polyacrylamide gel electrophoresis (SDSPAGE) 148, 150 Sodium hypochlorite 60 Soft Tick 270 Southern blotting 149 Specific soluble substance 36 Spore of Clostridium tetani 56 Spore Staining 37 Learning Objectives 37 Introduction 37 Principle 37 Requirements 37 Equipments 37 Reagents and lab wares 37 Preparation of malachite green stain 37 Preparation of safranine stain 37 Specimen 38 Procedure 38 Quality Control 38 Observations 38 Results and Interpretation 38 Viva 38 Key Facts 38

311

Spores of B. stearothermophilus 57 Spores of Bacillus stearothermophilus 56 Spores of Microsporidia 199 Sporothrix schenckeii 6 Spread plate inoculation procedure 14 Standard test of syphilis 125 Staphylococcus aureus 6, 166, 278 Staphylococcus aureus on nutrient agar 278 Sterile Mueller Hinton broth 97 Sterilisation 56, 60 Sterilization of Commonly Used Culture Media 56 Learning Objectives 56 Introduction 56 Principle 56 Sterilization by heat 56 Sterilization by filtration 56 Requirements 56 Equipments 56 Reagents 56 Specimen 56 Procedure 56 Quality Control 57 Autoclave 57 Hot air oven 57 Filters 57 Observations 57 Results and Interpretation 57 Key Facts 58 Viva 58 Sterilization of culture media 56 Stokes Method 95, 96 Learning Objectives 95 Introduction 95 Principle 95 Requirements 95 Equipments 95 Reagents and lab wares 95 Specimen 95 Procedure 95 Quality Control 95 Observations 96 Results and Interpretation 96 Key Facts 96 Viva 96 Streak plate inoculation procedure 14 Streptococcus MG agglutination 120 Streptococcus pneumoniae 169, 170, 261 Streptococcus pyogenes 172, 278 Streptococcus pyogenes on blood agar 278 Streptolysin O 121 Streptolysin S 121, 122 Streptomycin-resistant mutants 152 String test 183 Stuart’s broth 72 Subcutaneous mycoses 212 Suckling mouse 259, 258 Sulphur indole motility medium 86 Superficial mycoses 212 Swarming 11 Systemic mycoses 212

T TAB vaccine 117 Table 1-1 Types of condensers 4 Table 13-1 List of different type of media commonly used forisolation of bacteria 46 Table 14-1 Examples of bacteria showing different temperaturesfor their growth 48

312

Index

Table 16-1 Examples of bacteria grouped depending on theirrequirement of oxygen 52 Table 18-1 Different methods of sterilization of varioussubstances 58 Table 19-1 List of commonly used disinfectants and antiseptics 60 Table 20-1 Catalase positive and negative bacteria 63 Table 21-1 List of oxidase positive and negative bacteria 66 Table 22-1 List of coagulase positive bacteria 69 Table 23-1 Urease producing bacteria and fungi 72 Table 24-1 List of indole positive and negative bacteria 75 Table 25-1 List of MR positive and negative bacteria 77 Table 26-1 VP positive and negative bacteria 79 Table 27-1 Differences between Simmon’s citrate and Koser’scitrate 80 Table 27-2 List of citrate positive and negative bacteria 81 Table 29-1 List of H2S positive bacteria 86 Table 32-1 Comparison of Kirby-Bauer and Stokes methods 96 Table 34-1 Preparation of stock dilutions of the antibiotic stocksolutions 101 Table 4-1 Motile and non-motile bacteria 12 Table 43-1 Advantages and disadvantages of VDRL test 124 Table 44-1 Uses of gel diffusion tests 127 Table 48-1 Advantages and disadvantages of the immunofluorescencetest 136 Table 49-1 Enzymes and substrates used in the ELISA 140 Table 49-2 Advantages and disadvantages of ELISA test 140 Table 51-1 List of other types of electrophoresis in gels 149 Table 51-2 Calculation for X% separating or stacking gel 150 Table 52-1 Different types of mutations and their role inmicrobial infections 153 Table 53-1 Differences between F+ strain and Hfr strain 156 Table 57-1 Laboratory tests for differentiation of staphylococcalspecies 167 Table 58-1 Differences between Streptococcus pneumoniaeand Streptococcus mitis 170 Table 62-1 Differences between V. cholerae biotypeclassical and V. cholerae biotype El Tor 182 Table 65-1 List of bile-stained and non-bile stained eggs 193 Table 69-1 Other floatation methods of concentration stool 206 Table 7-1 List of Gram positive and Gram negative bacteria 24 Table 71-1 List of media used for fungal culture 214 Table 77-1 Differences between germ tubes and pseudohyphae 226 Table 79-1 Preparation of Yeast Nitrogen Base 230 Table 8-1 List of acid-fast structures 29 Table 82-1 The cell lines and indications for the viruses andcytopathic effects they produce 241 Table 83-1 The routes of inoculation of the egg and the virusesisolated 244 Table 84-1 Grades of the Quality of Drinking Water Supplies Determined by results of Periodic Escherichia coli andColiform counts 250 Table 85-1 List of bacteria that can be found in contaminatedmilk 253 Table 86-1 List of bacteria commonly found in air 254 Table 89-1 Laboratory animals and their usage 265 Table Applications of various tests used in a microbiologylaboratory 106 Tachyzoites 290 Taenia saginata 6 Taenia soluim 6 Tautomerization 154 TCBS 181 Tease mount preparation 217 Temperature Requirement for Growth of Bacteria 47 Learning Objectives 47 Introduction 47 Principle 47 Psychrophilic bacteria 47 Mesophilic bacteria 47 Thermophilic bacteria 47 Requirements 47 Equipments 47 Reagents and media 47

Specimen 47 Procedure 47 Quality Control 47 Observations 48 Results and Interpretation 48 Key Facts 48 Viva 48 Test for coliform bacilli 252 Test for pathogenic bacteria 249 Tetramethyl paraphenylene diamine dihydrochloride 160 Thick blood smear 201, 202, 204 Thin blood films 201, 202, 204 Thioglycollate broth 52 Thioglycollate broth culture 53 Tick borne encephalitis 271 Tick paralysis 271 Ticks 270 Toluidine blue 31 Total coliform count 248 Toxoplasma gondii 199, 258, 290 Trachoma 269 Transcription 144, 156 Transformation 156 Transversions 154 Trench fever 270 Treponema pallidum 287 Trichinella spiralis 6 Trichomonas vaginalis 8 Trichophyton verrucosum 236 Trichophyton violaceum 236 Triple sugar iron (TSI) agar 82 Triple sugar iron agar (TSI) test 82 Learning Objectives 82 Introduction 82 Principle 82 Requirements 82 Equipments 82 Reagents and lab wares 82 Specimen 82 Procedure 83 Quality Control 83 Observations 83 Results and Interpretation 83 Viva 83 Key Facts 84 Trombicula akamush 269 Trombicula dilensis 269 Trombiculid Mite 269 Tryptone broth 74 TSI agar 82, 84, 85, 86, 282 Tube agglutination tests 106 Tube coagulase test 69, 70 Tube dilution method 100 Tuberculin test 284 Tuberculous meningitis 29 Turbidity test 252 Tyndallization 58

U Universal Container 283 Universal donors 110 Urea 71, 227 Urea hydrolysis test 176 Urease 71, 72, 227, 228 Urease negative bacteria 72 Urease negative fungi 227 Urease positive bacteria 72 Urease positive fungi 227

Textbook of Practical Microbiology Urease producing bacteria 72 Urease producing fungi 72 Urease test 71, 72, 185, 227, 228, 281 Learning Objectives 71 Introduction 71 Principle 71 Requirements 71 Equipments 71 Reagents and glass wares 71 Specimen 71 Procedure 71 Quality Control 72 Positive control 72 Negative control 72 Observation 72 Results and Interpretation 72 Key Facts 72 Viva 72 Urease Test for fungi 227, 279 Learning Objectives 227 Introduction 227 Principle 227 Requirements 227 Equipments 227 Reagents and lab wares 227 Specimen 227 Procedure 227 Quality Control 227 Observations 227 Results and Interpretation 227 Key Facts 228 Viva 228

V VDRL test 123, 125 Learning Objectives 123 Introduction 123 Principle 123 Requirements 123 Equipments 123 Reagents and glass wares 123 Preparation of buffered saline solution 123 Preparation of un buffered saline solution 123 Specimen 123 Procedure 123 Preparation of serum 123 Preparation of antigen emulsion 124 Maturation of the antigen 124 Qualitative serum test 124 Quantitative serum test 124 Quality control 124 Observations 124 Results and Interpretation 124 Qualitative serum test 124 Quantitative serum test 124 Key Facts 125 Viva 125 VDRL-ELISA 123, 125 Veronal buffer 130 Viable count 252 Vibrio cholerae 181, 287 Vibriostatic (O/129) agent 182 Voges-Proskauer (VP) test 182 Voges-Proskauer Test 78 Learning Objectives 78 Introduction 78 Principle 78 Requirements 78

Equipments 78 Reagents and lab wares 78 Specimen 78 Procedure 78 Quality Control 79 Positive control 79 Negative control 79 Observations 79 Results and Interpretation 79 Key Facts 79 Viva 79 Volutin granules 31 VP broth 78 VP negative bacteria 79 VP positive bacteria 79 VP test 79

W Water fleas 271 Weil-Felix test 119 Learning Objectives 119 Introduction 119 Principle 119 Requirements 119 Equipments 119 Reagents and glass wares 119 Specimen 119 Procedure 119 Preparation of master dilution of the serum 119 Performance of the test 119 Quality Control 120 Observations 120 Results and Interpretation 120 Key Facts 120 Viva 120 West Nile fever 269 Western blot 150 Wet mount preparation 191, 206 Widal test 116, 117 Learning Objectives 116 Introduction 116 Principle 116 Requirements 116 Equipments 116 Reagents and glass wares 116 Specimen 116 Procedure 116 Preparation of master dilution of the serum 116 Performance of the test 116 Quality Control 117 Observations 117 Results and Interpretation 117 Viva 117 Key Facts 118 Wright’s stain 201 Wuchereria bancrofti 291 Wuchereria bancrofti microfilaria 291

X Xylose lysine deoxycholate agar 86

Y Yaws 269 Yeast-like fungi 229 Yellow fever 269 Yolk sac 243 Yolk sac 244

313

314

Index

Z Zephiran 60 Ziehl Neelsen staining 288 Ziehl-Neelsen acid-fast staining 27 Ziehl-Neelsen stain 29, 37

Zinc dusts 89 Zinc sulphate floatation method 206 Zone reactions 124, 125 Zygomycetes 6

1 / Running Head

TEXTBOOK OF PRACTICAL MICROBIOLOGY

The intent of the book is to provide recent information and explain in detail the routine diagnostic methods performed in a Microbiology laboratory. Every effort has been made to incorporate all aspects of practical microbiology. A sincere effort is made to provide the essential underlying principles of practical microbiology, to help students to perform various practicals, and to learn and apply the knowledge of practical microbiology in clinical medicine

Subhash Chandra Parija is Director-Professor and Head of Microbiology at JIPMER, Pondicherry.

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Ahuja Publishers New Delhi

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