Practical Manual of Medical Microbiology-book53.pdf

Practical Manual of Medical Microbiology (For Medical, Dental and Paramedical Students)

Practical Manual of Medical Microbiology (For Medical, Dental and Paramedical Students)

CP Prince MSc (Medical Microbiology), PhD, FAGE

Lecturer Department of Microbiology Mother Theresa Institute of Health Sciences (A Government of Puducherry Institution) Puducherry, India

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Published by Jitendar P Vij Jaypee Brothers Medical Publishers (P) Ltd Corporate Office 4838/24 Ansari Road, Daryaganj, New Delhi - 110 002, India, +91-11-43574357 (30 lines) Registered Office B-3 EMCA House, 23/23B Ansari Road, Daryaganj, New Delhi 110 002, India Phones: +91-11-23272143, +91-11-23272703, +91-11-23282021, +91-11-23245672, Rel: +91-11-32558559 Fax: +91-11-23276490, +91-11-23245683 e-mail: [email protected], Website: www.jaypeebrothers.com Branches • 2/B, Akruti Society, Jodhpur Gam Road Satellite Ahmedabad 380 015 Phones: +91-79-26926233, Rel: +91-79-32988717 Fax: +91-79-26927094 e-mail: [email protected] • 202 Batavia Chambers, 8 Kumara Krupa Road, Kumara Park East Bengaluru 560 001 Phones: +91-80-22285971, +91-80-22382956, +91-80-22372664 Rel: +91-80-32714073, Fax: +91-80-22281761 e-mail: [email protected] • 282 IIIrd Floor, Khaleel Shirazi Estate, Fountain Plaza, Pantheon Road Chennai 600 008 Phones: +91-44-28193265, +91-44-28194897, Rel: +91-44-32972089 Fax: +91-44-28193231 e-mail: [email protected] • 4-2-1067/1-3, 1st Floor, Balaji Building, Ramkote Cross Road Hyderabad 500 095 Phones: +91-40-66610020, +91-40-24758498, Rel:+91-40-32940929 Fax:+91-40-24758499 e-mail: [email protected] • No. 41/3098, B & B1, Kuruvi Building, St. Vincent Road Kochi 682 018, Kerala Phones: +91-484-4036109, +91-484-2395739, +91-484-2395740 e-mail: [email protected] • 1-A Indian Mirror Street, Wellington Square Kolkata 700 013 Phones: +91-33-22651926, +91-33-22276404, +91-33-22276415 Rel: +91-33-32901926, Fax: +91-33-22656075, e-mail: [email protected] • Lekhraj Market III, B-2, Sector-4, Faizabad Road, Indira Nagar Lucknow 226 016 Phones: +91-522-3040553, +91-522-3040554 e-mail: [email protected] • 106 Amit Industrial Estate, 61 Dr SS Rao Road, Near MGM Hospital, Parel Mumbai 400012 Phones: +91-22-24124863, +91-22-24104532, Rel: +91-22-32926896 Fax: +91-22-24160828 e-mail: [email protected] • “KAMALPUSHPA” 38, Reshimbag, Opp. Mohota Science College, Umred Road Nagpur 440 009 (MS) Phone: Rel: +91-712-3245220, Fax: +91-712-2704275 e-mail: [email protected] USA Office 1745, Pheasant Run Drive, Maryland Heights (Missouri), MO 63043, USA Ph: 001-636-6279734 e-mail: [email protected], [email protected] Practical Manual of Medical Microbiology © 2009, Jaypee Brothers Medical Publishers All rights reserved. No part of this publication should be reproduced, stored in a retrieval system, or transmitted in any form or by any means: electronic, mechanical, photocopying, recording, or otherwise, without the prior written permission of the author and the publisher. This book has been published in good faith that the material provided by author is original. Every effort is made to ensure accuracy of material, but the publisher, printer and author will not be held responsible for any inadvertent error(s). In case of any dispute, all legal matters are to be settled under Delhi jurisdiction only. First Edition: 2009 ISBN 978-81-8448-637-7 Typeset at

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PREFACE

Practical Manual of Medical Microbiology is aimed to help the teachers and students to conduct practical/demonstration classes and to solve the difficulty of maintaining the practical record book. A sincere effort is made to provide brief knowledge on the principles and procedures of common laboratory experiments. The figures and photographs especially the line diagrams illustrated in this book will be useful to perform various experiments and write the observations and reports. The list of spotters and identification points given in the last chapter will help the students to excel in their practical examinations. The content of the book covers the syllabus requirements of many Universities and other regulatory bodies like Medical, Dental and Nursing Councils. The WHO recommended test procedures and quality assurance programmes which are mentioned in this book. These procedures and programmes may be helpful to standardise and streamline the experiments in newly established medical institutions all over the world. This book is tailored to the need of students of MBBS, BDS, BSc (MLT), BSc (Microbiology), DMLT and other paramedical courses and those who work in the field of microbiology needing short and concise information. I am extremely grateful to Dr V Balu, Dean, Mother Theresa Institute of Health Sciences (MTIHS), Dr V Gopal, Principal, College of Pharmacy, MTIHS and Dr Helen PS Mannuel, former DirectorProfessor, Madras Medical College for their constant encouragement and support. I owe special thanks and gratitude to my colleagues and family for their support and help. Readers’ suggestions and comments will help for further improvement of the book in future editions. CP Prince

CODE OF PROFESSIONAL CONDUCT FOR MEDICAL LABORATORY PERSONNEL

1. Place the well-being and service of the sick above your own interests. 2. Be loyal to your medical laboratory profession by maintaining high standards of work and striving to improve your professional skills and knowledge. 3. Work scientifically and with complete honesty. 4. Do not misuse your professional skills or knowledge for personal gain. 5. Never take anything from your place of work that does not belong to you. 6. Do not disclose to a patient or any unauthorised person the results of your investigations. 7. Treat with strict confidentiality any personal information that you may learn about a patient. 8. Respect and work in harmony with the other members of your hospital staff or health centre team. 9. Be, at all times, courteous, patient, and considerate to the sick and their relatives. 10. Promote health care and the prevention and control of disease. 11. Follow safety procedures and know how to apply First Aid. 12. Do not drink alcohol during laboratory working hours or when on emergency stand-by. 13. Use equipment and laboratory ware correctly and with care. 14. Do not waste reagents or other laboratory supplies. 15. Fulfill reliably and completely the terms and conditions of your employment.

CONTENTS 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.

Laboratory Safety ............................ 1 First Aid .................................. 10 Hand Washing ............................. 12 Units .................................... 14 Microscope ................................ 17 Micrometry ................................ 24 Sterilisation ................................ 27 Hanging Drop Preparation ..................... 37 Preparation and Fixation of Smears ............... 41 Methylene Blue Staining ....................... 45 Indian Ink Staining .......................... 47 Gram’s Stain ............................... 49 Ziehl-Neelsen’s Stain (Acid Fast Stain) ............ 53 Albert’s Stain .............................. 56 Leishman’s Stain ............................ 59 Preparation and Cleaning of Glassware ............ 62 pH in Microbiology .......................... 67 Bacteriological Media ......................... 69 Inoculation of Culture Media ................... 77 Anaerobic Cultivation ........................ 84 Important Bacterial Pathogens and the Diseases ...... 88 Collection of Clinical Materials .................. 95 for Microbiological Investigations Biochemical Tests and Identification of Bacteria ..... 106 O/F Test (Hugh and Leifson’s Test) .............. 108 Catalase Test .............................. 110 Oxidase Test .............................. 112 Sugar Fermentation Test ...................... 115 Nitrate Reduction Test ....................... 118 Hydrogen Sulphide Production Test ............. 120

x

Practical Manual of Medical Microbiology 30. 31. 32. 33. 34. 35. 36. 37. 38. 39. 40. 41. 42. 43. 44. 45. 46. 47. 48. 49. 50. 51. 52. 53. 54. 55. 56. 57. 58. 59. 60.

Urease Test ............................... 122 Citrate Utilisation Test ....................... 124 Voges–Proskauer Test (VP Test) ................ 126 Methyl Red Test (MR Test) .................... 128 Indole Test ............................... 130 Bile Solubility Test .......................... 133 Coagulase Test ............................ 135 Antimicrobial Susceptibility Testing ............. 138 Brucella Agglutination Test ................... 148 Anti-Streptolysin O (ASO) Test ................. 150 CRP Screen Latex Agglutination Slide Test ......... 152 VDRL Test ............................... 156 Treponema Pallidum Haemagglutination Assay .... 159 (TPHA) Widal Test ............................... 162 Enzyme Linked Immunosorbent Assay (ELISA) ..... 165 Experimental Animals ....................... 172 Potassium Hydroxide Wet Mount ............... 175 Lactophenol Cotton Blue Mount ................ 177 Culture of Fungi ........................... 179 Fungal Slide Culture (Riddle’s Method) ........... 182 Identification of Fungal Isolates ................ 184 Germ Tube Test ............................ 195 Diagnosis of Virus Infections .................. 197 Lab Diagnosis of Malaria ..................... 201 Parasitological Examination of Faeces ............ 210 Medical Entomology ........................ 220 Bacteriological Examination of Water ............ 233 Microbiology of Milk ........................ 239 Lab Diagnosis of Tuberculosis ................. 241 Urinary Tract Infection ....................... 247 Spotters ................................. 253

Index .................................... 267

Laboratory Safety 1

1 Laboratory Safety ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Laboratory safety is a vital component of functioning of any laboratory. Safety procedures and precautions to be followed in the Microbiology laboratory should be designed to: • Restrict microorganisms present in specimens or cultures to the vessels in which they are contained. • Prevent environmental microorganisms (normally present on hand, hair, clothing, laboratory benches or in the air) from entering specimens or cultures and interfering with the results of the studies. Laboratory Biosafety Levels Four Biosafety levels have been recommended based on the type of microbes you are working with. Biosafety Level -1 (BSL-1): Adherence to standard microbiological practices. No special requirement as regards containment equipment. Biosafety Level-2 (BSL-2): In addition to the use of standard microbiological practice, laboratory coats, decontamination of infectious wastes, limited access, protective gloves and display of biohazard sign and partial containment equipment are the requirements for this level. Most peripheral and intermediate laboratories need BSL-1 or BSL-2 laboratory facilities.

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BSL-3: In addition to BSL-2, it has special laboratory clothing, controlled access to laboratory and partial containment equipment. BSL-4: BSL-3 plus entrance through change room where laboratory clothing is put on, shower on exit, all wastes are decontaminated before exit from the facility. It requires maximum containment equipment. Laboratory facilities in BSL-2 (Figs 1.1 and 1.2) • Laboratory should be designed in such a way that it can be easily cleaned. • Laboratory contains a sink for washing. • Laboratory tops are impervious to water but resistant to acids, alkalis and organic solvents. • An autoclave to decontaminate infectious material is available. • Illumination is adequate for all laboratory activities. • Storage space is adequate. • Preventive measures against laboratory infections These are aimed to protect workers, patients and cultures. Following steps are suggested: • Perform adequate sterilization before washing or disposing waste. • Provide receptacle for contaminated glassware. • Provide safety hood. • Ensure that tissues are handled and disposed of properly. • Promote regular hand washing and cleaning of bench tops. • Ensure use of gloves. • Provide mechanical pipetting devices. • Protect patients from laboratory personnel with skin or upper respiratory tract infections. • Provide special disposal containers for needles and lancets. Pipetting Pipetting and suctioning have been identified as the significant and consistent causes of occupational infections. Various important precautions that must be taken while pipetting are: • Develop pipetting techniques that reduce the potential for creating aerosols.

Laboratory Safety 3

Figure 1.1: Hazard warning symbols

Figure 1.2: Biological safety cabinets

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• • • • • •

Plug pipettes with cotton. Avoid rapid mixing of liquids by alternate suction and expulsion. Do not forcibly expel material from a pipette. Do not bubble air through liquids with a pipette. Prefer pipettes that do not require expulsion of last drop of liquid. Drop material having pathogenic organisms as close as possible to the fluid or agar level. • Place contaminated pipettes in a container having suitable disinfectant for complete immersion. A variety of pipettes are available. Selection should depend upon the ease of operation and the type of work to be performed. Hypodermic Syringes and Needles Accidents involving the use of syringes and needles while drawing blood from patients or performing experiments on laboratory animals are among the most common causes of occupational infections in laboratories and health care facilities. They account for almost 25% of the laboratory-acquired infections that occur by accidents. The practices which are recommended for hypodermic needle and syringes are: • Avoid quick and unnecessary movements of the hand holding the syringe. • Examine glass syringes for chips and cracks, and examine needles for barbs and plugs. • Use needle locking (Luer Lock type) syringes only and be sure that needle is locked securely. • Wear surgical or other gloves. • Fill syringes carefully to minimize air bubbles and frothing. • Expel excess air, liquid and bubbles vertically into a cotton pledget moistened with suitable disinfectant. • Do not use syringe to forcefully expel infectious fluid into an open vial for mixing. Mixing with a syringe is appropriate only if the tip of the needle is held below the surface of the fluid in the tube. • Do not bend, shear, recap or remove the needle from syringe by hand.

Laboratory Safety 5 • Place used needle-syringe units directly into a puncture-resistant container and decontaminate before disassembly, reuse or disposal. Opening Containers The opening of vials, flasks, petri dishes, culture tubes, embryonated eggs, and other containers of potentially infectious materials poses potential but subtle risks of creating droplets, aerosols or contamination of the skin or the immediate work area. The most common opening activity in most health care laboratories is the removal of stoppers from containers of clinical materials. It is imperative that specimens should be received and opened only by personnel who are knowledgeable about occupational infection risks. Various precautions that can be taken in this regard are: • Open containers with clinical specimens in well-lighted and designated areas only. • Wear a laboratory coat and suitable gloves. • If possible, use a plastic-backed absorbent paper towel to: – Facilitate clean-up – Reduce generation of aerosols • Specimens which are leaking or broken may be opened only in safety cabinets. Tubes containing bacterial cultures should be handled with care. Vigorous shaking of liquid cultures creates a heavy aerosol. When a sealed ampoule containing a lyophilized or liquid culture is opened, an aerosol may be created. Ampoules should be opened in a safety cabinet(Fig.1.3). Laboratory Access • As far as possible children and pregnant women visitors should not enter the microbiological laboratories. • Appropriate signs should be located at points of access to laboratory areas directing all visitors to a receptionist or receiving office for access procedures. • The universal biohazard symbol shall be displayed at specific laboratories in which manipulations of organisms with moderate

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Figure 1.3: Biological safety cabinet

and heavy risk are being carried out. Only authorized visitors shall enter the laboratory showing universal biohazard sign (Fig. 1.4). Doors displaying biohazard symbol shall not be propped open, but shall remain closed when in use.

. Figure 1.4: Universal biohazard sign

Laboratory Safety 7 Clothing • All employees and visitors in microbiological laboratories shall wear laboratory clothing and laboratory shoes or shoe covers. • Disposable gloves shall be worn wherever radiological, chemical, carcinogenic materials or virus preparations of moderate to high risk are handled. • Laboratory clothing including shoes shall not be worn outside the work area. Accidents in Laboratory In the microbiological laboratory, infections pose the most frequent risk. The important pathogens are: Hepatitis B virus, Shigella spp. HIV, Salmonella spp. including S typhi Brucella spp. Bacillus anthracis Leptospires Yersinia pestis Mycobacteria spp. Histoplasma Accidents and Spills The order of priorities is as follows: • Protection of personnel • Confinement of contamination • Decontamination of personnel • Decontamination of area involved

Decontamination of skin: The area is washed thoroughly with soap and water. Detergents or abrasive materials must not be used and care must be taken not to damage the skin. Decontamination of cuts\eyes: These are irrigated with water taking care to prevent the spread of contamination from one area to another. Decontamination of clothing: Contaminated garments should be removed immediately and placed in a container. They should not be removed from the spill location until contamination has been monitored.

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Decontamination of work surfaces • Flood the total spillage area including the broken container with disinfectant. • Leave undisturbed for 10 minutes. • Mop with cotton wool or absorbent paper. • Wear disposable gloves, apron and goggles. • If a dustpan and brush or forceps have been used these too require disinfection. • For blood or viruses, hypochlorites (10 gm/L) are used. • Do not use hypochorite solution in centrifuges. • Use activated gluteraldehyde (20 gm/L) on surfaces for viral decontamination. • Place all potentially contaminated materials in a separate container and retain until monitored. • Restrict the entry to such an area until contamination monitoring has been carried out. Management of Laboratory Accidents An adequately equipped first-aid box should be kept in the laboratory in a place that is known and accessible to all members of staff. The box must be clearly marked and preferably be made of metal or plastic to prevent from damage by pests. A medical officer should be consulted regarding the contents of the box. A first-aid chart giving the immediate treatment of cuts, burns, poisoning, shock and collapse, should be prepared and displayed in the laboratory. General Laboratory Directions for Safety The salient general laboratory directions which must be obeyed by allare: • Long hair should be bound back neatly away from shoulders. • Do not wear any jewellery to laboratory sessions. • Keep fingers, pencils, bacteriological loops, etc. out of your mouth. • Do not smoke in the laboratory. • Do not lick labels with tongue (use tap water). • Do not drink from laboratory glassware.

Laboratory Safety 9 • Do not wander about the laboratory; uncontrolled activities cause: – Accidents – Distract others – Promote contamination • Do not place contaminated pipettes on the bench top. • Do not discard contaminated cultures, glassware, pipettes, tubes or slides in wastepaper basket or garbage can. • Avoid dispersal of infectious materials. • Operate centrifuges, homogenizer and shakers safely. • Immunize the laboratory workers against vaccine-preventable diseases such as hepatitis B, meningococcal meningitis, rabies, etc.

2 First Aid ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Knowledge of first aid can help to reduce suffering and consequences of serious accidents. In some situations, first aid can be life saving. All laboratory workers should receive a basic practical training in First Aid, with particular attention being paid to the types of accidents which may occur in the laboratory. First Aid Box An adequately equipped first aid box should be kept in the laboratory. The box should be clearly marked with a red cross. The common items kept in a First Aid Box are given below. The laboratory incharge should regularly check the items; replace the missing items or medicines that may have expired. 1. First aid manual 2. Sterile gauze 3. Adhesive tape 4. Adhesive bandages in several sizes 5. Elastic bandage 6. Antiseptic wipes 7. Soap 8. Antibiotic cream (triple antibiotic ointment) 9. Antiseptic solution (like hydrogen peroxide)

First Aid 11 10. 11. 12. 13. 14. 15. 16. 17. 18. 19. 20. 21. 22. 23.

Hydrocortisone cream (H) Acetaminophen and ibuprofen Tweeters Sharp scissors Safety pins Disposable instant cold packs. Calamine lotion Alcohol wipes and ethyl alcohol Thermometer Plastic gloves (at least two pairs) Flash light and extra batteries Mouth piece for administering CPR List of emergency phone numbers Blanket (stored nearby)

3 Hand Washing ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION “Soap and common sense can prevent 80% of Nosocomial of infections”. This fact points to the importance of soap and hand washing in the control of infectious diseases. Hand hygiene has been described as the single most effective means of preventing spread of infections. The aim of hand washing is to remove transient microorganisms (not commensal organisms) as soon as possible following acquisition from contact with possible sources, or immediately before performing invasive procedures and touching susceptible patients or susceptible sites. Method Proper hand washing can easily achieve by thorough hand washing technique with soap and water using mechanical friction for 10-15 seconds followed by drying. Surgical hand wash involves washing both hands and fore arms using a defined technique for at least 2 minutes (Fig. 3.1). In order to disinfect clean hands, alcohol rub can be applied following washing. Alternatively, alcohol based product can be used which are shown to be very effective when running water, soap and

Hand Washing 13

Figure 3.1: Hand washing procedure

towels are not available. Alcohol would not be effective for soiled hands which require prior washing. In hospitals and laboratories hand washing facilities should be conveniently located in sufficient numbers. The facility should include sink of adequate size with elbow or sensor operated tap and wall mounted liquid hand washing agent like soap or disinfectant and paper towel unit. Waste bin with foot-operated lid should be provided along side. Commonly used disinfectant solutions are chlorhexidine, povidone iodine and commercially available antiseptic solutions like Dettol or Savlon.

4 Units ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

The international system of units has been developed and agreed internationally in the interest of world health. It overcomes language barriers, enabling an exchange of health information with in a country and between nations. SI units (Systeme International d’ Unites) are commonly used. There are seven basic SI units, meter, kilogram, second, mole, ampere, Kelvin and candela. All other units are derived from these seven base units. Some SI derived units have been given special names. Sl.No. SI base units 1. 2. 3. 4. 5. 6. 7.

Metre Kilogram Second Mole Ampere Kelvin Candela

Symbol

Quantity measured

m kg S mol A K cd

length mass time amount of substance Electric current temperature luminous intensity.

Units 15

Sl.No. SI derived units

Symbol 2

1. Square metre m 2. Cubic metre m3 3. Metre per second m / s Sl.No. Named derived unit 1. 2. 3. 4. 5. 6. 7.

Hertz Joule Newton Pascal Watt Volt Degree

Quantity measured area volume speed

Symbol

Quantity measured

Hz J N Pa W V o C

frequency energy, quantity of heat Force Pressure Power Electric potential difference Celsius temperature

SI Unit Prefixes To enable the measurement of larger or smaller units, SI system also includes a set of prefixes. The use of a prefix makes a unit larger or smaller. Example: if the prefix milli is put in front of the metere (millimeter) this indicates that the unit should be divided by a thousand (10–3) Prefix Deci Centi Milli Micro Nano Pico Femto

Symbol d C M u n P f

Function –1

10 10 –2 10 –3 10 –6 10 –9 10 –12 10 –15

Divided by 10 100 1000 1000 000 1000 000 000 1000 000 000 000 1000 000 000 000 000

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Practical Manual of Medical Microbiology

Prefix Deca Hecta Kilo Mega Giga Tera Peta

Symbol da h K M G T P

Function 1

10 10 2 10 3 10 6 10 9 10 12 10 15

Multiplied by 10 100 1000 1000 000 1000 000 000 1000 000 000 000 1000 000 000 000 000

5 Microscope ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Microscope was invented by Antony Van Leuwenhoek (1632-1723). He was a Dutch lens maker and was the first person to observe Bacteria. Microscope is an essential optical instrument of Microbiology laboratory. It consists of combination of lenses which will give a magnified image of minute objects or micro organisms like Bacteria, Fungi, and Protozoa. Types of Microscopes 1. 2. 3. 4. 5. 6. 7. 8. 9.

Simple microscope Compound microscope Ultraviolet microscope Fluorescent microscope Polarizing microscope Dark ground microscope Phase contrast microscope Inverted microscope Electron microscope

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Microscope that is suitable for the study of microorganisms is the light compound Microscope. This consists of two converging lenses fixed at the ends of a brass tube. The lens which is nearer to the object is called “Objective” and the lens which is close to the eye is called Eyepiece or “Ocular”. The final image can be observed through the ocular. The objective magnifies the specimen to a definite amount and produces a real, inverted intermediate image of the object, which lies within the principle focus of the eyepiece. The eyepiece further magnifies the image formed by the objective so that the image seen by the eye has a magnification, equal to the product of magnification of the two systems. The individual magnification of objectives and eyepiece are engraved on each part. The final image seen is thus inverted, magnified and virtual. Parts of Compound Microscope Based on the manufacturer the parts and adjustments may slightly vary from each other. Microscopes with fixed stage and mechanical stage are available. Microscope with mechanical stage is more convenient for microbiological studies (Fig. 5.1). I Mechanical parts Base (Foot) Limb (Arm) Stage Adjustment knobs Revolving nose piece II Magnifying parts Objective Ocular III Illuminating parts Sub stage condenser Iris diaphragm Filter holder Mirror Electric bulb MECHANICAL PARTS a. Base: It forms the stand or foot of the microscope, often horse shoe shaped to give the stability. The mirror or electric bulb illuminator is attached to the base.

Microscope 19

Eye piece Body tube Body

Nose piece Limb Objective Stage Condenser Coarse adjustment Mirror

Fine adjustment

Base

Figure 5.1: Compound microscope

b. Limb: It forms the arm which bears the illuminating parts, stage and the observation tube. In some microscopes the limb is attached to the foot by a hinged joint so that the microscope can be set at a comfortable angle for the observer. c. Stage: The stage is a plat form which accommodates a glass slide on which the object to be examined is mounted. It has an aperture in its centre to permit light to reach the object. The stage can be of 2 types: i. A fixed stage on which the object is fixed by clips. ii. The mechanical stage holds the slide secure and allows the specimen to be moved smoothly backwards, forwards or sideways. Sometimes a scale is fitted to two sides of the stage to show the extent of the movement. This is called the Vernier Scale and it is useful to trace a part of the blood film or sputum smear that you need to re-examine or show to your supervisor. d. Adjustment knobs: They are used to focus the object two types of knobs will be present (1) Coarse adjustment for initial adjustment and (2) Fine adjustment for getting a clear image.

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In some new microscopes focusing is done by moving the stage (movable stage) by coarse and fine adjustments. In older versions, the stage is fixed and focusing is done by moving the Body tube by coarse and fine adjustment knobs. e. Revolving nose piece: A number of objective lenses of different magnifications are screwed to the nosepiece of the microscope, which can be revolved to increase or decrease the magnification of the specimen being examined by selecting the objective lens. MAGNIFYING PARTS This consists of eyepieces and objectives. They are kept separated in a graduated tube. Objectives are referred to by their magnifying power, which is marked on the side. The microscope commonly used in student laboratories will have the following objectives (Fig. 5.2).

Figure 5.2: Objectives of microscope

10 × (Low power objective) 40 × (High power objective) 100 × (Oil immersion objective) As the magnification differs between objectives, so does the working distance. The working distance is the distance between the front lens of the objective and the specimen on the stage (when the specimen is in focus). The higher the magnifying power of the objective the shorter is the working distance. Working distances for the standard objectives are likely to be indicated as follows.

Microscope 21 10 × – 15.98 mm 40 × – 4.31 mm 100 × – 1.81 mm (This gap is filled with cedar wood oil) The eyepiece/ocular commonly used will have a magnification of 10 ×. Oculars with 5 ×, 6 ×, 15 × and 20 × are also available. ILLUMINATING PARTS Sub-stage Condensers It is made up of a system of convex lenses, which focus light from the illuminating source on the place of the object. The height of the condenser can be varied by a rack and pinion mechanism. Lowering of condenser diminishes illumination whereas raising the condenser increases the illumination. While using oil immersion objective, the condenser is completely raised as it requires more light. When the other objectives are used, it is to be lowered suitably. Iris Diaphragm Immediately below the condenser and incorporated in the same mount is the sub stage iris diaphragm operated by a small lever which protrudes to one side. Opening or closing of this iris diaphragm controls the amount of light reaching the object. Iris diaphragm is opened widely when the oil-immersion objective is used as it requires maximum light and closed partially when the other objectives are in use. Filter Holder Just below the iris diaphragm is ring shaped filter holder designated to carry circular, blue colored glass filters required to reduce the excessive red or yellow component of some light sources. Mirror Fitted to the base, below the condenser is a Plano- concave mirror. This is the illuminating source. It helps to reflect the light to the sub stage condenser. The flat surface (Plane mirror) is used whenever the oil immersion objective is employed. The concave mirror is employed with low and high power objectives.

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Electric Bulb Instead of mirror in newer models there will be an electric bulb which will act as source of illumination. It can also be replaced by mirror. The intensity of light can also be adjusted by the regulator. MAGNIFICATION It is defined as the degree of enlargement of the image of the object achieved by the microscope. The total magnification achieved by various objectives are given in the Table 5.1. Table 5.1: Total magnification Objective

Ocular

Mirror

10 ×

10 ×

40 × 100 ×

Oil Condenser

Iris diaphragm

Magnification

Concave No Low

Closed

10 × 10 = 100 times

10 ×

Concave No Middle

Half Open

40 × 10 = 400 times

10 ×

Plane

Open

100 × 10 = 1000 times

Yes Raised

Adjustment of Microscope Adjust the various parts as follows. For unstained preparation: 1. Lower the condenser 2. Close the iris diaphragm (Fig. 5.3) 3. Use concave mirror 4. Focus under low power and then turn to high power. For Stained preparations: 1. Lower the condenser 2. Use concave mirror 3. Adjust the iris diaphragm to give an even illumination of field (Fig. 5.3). 4. Focus under low power and then turn to high power, if required. Figure 5.3: Adjustments of diaphragm

Microscope 23 For Oil-immersion Examination: 1. Raise the condenser completely. 2. Open the iris diaphragm 3. Use plane mirror 4. Place a drop of cedar wood oil on the object and focus under oil immersion lens. Rack oil immersion objective down till its tip dips in the oil. Using the fine adjustment focus the object. 5. After this, remove the oil from the objective and the object with lens cleaning paper. 6. Leave low power objective in position till further use. Note: Cedar wood Oil has same refractive index as that of glass, Addition of oil in the gap between objective and object prevents refraction of light rays in order to get a bright image of the object. Care of Microscope • Clean the microscope with a clean soft cloth • The Objectives and oculars must be cleaned with lens paper • Alcohols should not be used as it dissolves the cement, which binds the lenses. • Direct sunlight on to the mirror should be avoided. • For prolonged work, artificial light of particular wavelengths are advisable. • When not used, protect it from dust and damages.

6 Micrometry ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION The measurement of objects using a calibrated eye piece scale (micrometer) is micrometry. This is used to measure the size of objects observed under microscope. Measurement of size is useful for the identification of microorganism especially cyst and Ova of Parasites. Requirement 1. Eyepiece micrometer: A suitable line scale for the eye piece micrometer is divided into 50 divisions. 2. Stage Micrometer: A suitable Scale for the stage micrometer is one that measures 2 mm in length with each large division measuring 0.1 mm (100 μm) and each small division measuring 0.01 mm (10 μm) (Fig. 6.1). Method 1. Caliberation of eye piece micrometer 2. Measuring an object with calibrated eye piece micrometer.

Micrometry 25

Figure 6.1: Micrometry: Stage micrometer scale (upper) and ocular micrometer (lower)

Calibration of Eye Piece Micrometer The eyepiece micrometer will require calibration for each objective of the microscope at which measurements will be required. For parasitology, the scale should be calibrated for the 40 × objective and for Bacteriology it is useful to calibrate for the 100 × objective. A table can be prepared for each objective giving measurement 1 to 50 divisions of the eye piece Scale. 1. Remove the normal eyepiece and insert the eye piece micrometer in the tube of the microscope. 2. Place the stage micrometer slide on the stage of the microscope. 3. Focus the stage micrometer scale using the required objective. 4. Adjust the field until the 0 line of the eye piece Scale aligns exactly with the ‘0’ line of the stage. 5. Look along the Scales and note where a division of the eye piece scale aligns exactly with a division of stage scale.

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6. Measure the distance between ‘0 point and where the alignment occurs. The measurements Calibration Scale are 0.1mm to 2.0 mm each small division measures 0.01mm. 7. Count the number of divisions of the eye piece Scale covered between the ‘0’ point and where the alignment occurs. 8. Calculate the measurement of 1 of the divisions of the eye piece Scale in 1 μm.

Example: Distance measured = Number of divisions =

0.2 mm 27

0.2 = 0.0074 mm 27 To Convert mm to μm = 0.0074 × 1000 = 7.4 μm

1 division measures

=

Measuring the Object • Remove the stage micrometer • Place the object slide • With eye piece micrometer measure divisions covered for the object (e.g. Cyst or ova) • Refer the prepared table for the objective being used to obtain the measurement of size.

7 Sterilisation ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Sterilisation is defined as the destruction or removal of all microorganisms and their spores. Disinfection is the destruction of many microorganisms but not usually the bacterial spores. Sterilization is usually achieved with the help of heat whereas chemical agents are employed to effect disinfection. Sterilisation and disinfection are part of the daily routine of microbiological laboratories and constitute a vital activity which ensures that cultures, containers, media and equipment are treated in such a way that only the inoculated organisms will grow while all others will be eliminated. Sterilisation by Heat This can be achieved by autoclaving, by exposing articles to dry heat in hot air ovens or boiling. Autoclave Autoclaves can sterilise anything that can withstand a temperature of 121oC for 30 minutes. A pressure cooker used in homes for cooking purposes can also be used as a makeshift autoclave (Fig. 7.1).

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Figure 7.1: Steam circulation in a double jacketed autoclave

The containers having clinical material are subjected to heat treatment in the autoclave after which these are emptied and washed and put back into service. Only autoclaves designed for laboratory work and capable of dealing with a mixed load should be used. Porous load and bottle fluid sterilisers are rarely satisfactory for laboratory work. There are two varieties of laboratory autoclaves. 1. Pressure cooker type 2. Gravity displacement models with automatic air and condensate discharge. Pressure-cooker type Laboratory Autoclaves The most common type is a device for boiling water under pressure. It has a vertical metal chamber with a strong metal lid which can be fastened down and sealed with a rubber gasket. An air and steam discharge tap, pressure gauge and a safety valve are fitted in the lid. Water in the bottom of the autoclave is heated by external gas burners, an electric immersion heater or a steam coil (Fig. 7.2).

Sterilisation 29

Figure 7.2: Vertical autoclave

Operating Instructions • Ensure that there is sufficient water inside the chamber. • Load the autoclave and fasten the lid keeping the discharge tap open. • Adjust the safety valve to the required temperature and turn the heat on. • Allow the mixture of air and steam to pass out freely till all air has been discharged. • Close the air discharge tap and let the steam pressure rise within the chamber till it attains a temperature of 121oC (1.5 kg/cm2). • Hold on the pressure for 15 minutes. • Turn off the heat and let the autoclave cool. • Slowly open the air and steam discharge taps after the pressure gauge has reached zero.

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• Allow the material to cool before these are handled (usually agar bottles take hours before these become safe to handle).

Autoclave with air discharge by gravity displacement These are usually rectangular in shape and arranged horizontally. These autoclaves have a jacket around the chamber (Figs 7.1 and 7.3).

Figure 7.3: Horizontal autoclave

Figure 7.4: Candle filter

Sterilisation 31 Operating Instructions • Bring the jacket of the autoclave to operating temperature. • Load the chamber, close the door and open the steam valve so that steam can freely enter the top of the chamber. Air and condensate shall automatically flow out through the drain at the bottom (Fig. 7.4). • When the drain thermometer reaches the required temperature, allow further period for the load to reach that temperature (this has to be determined initially and periodically for each autoclave). • Continue the autoclave cycle for the holding period. • Close the steam valve and let the autoclave cool till a temperature of 80oC is reached. • Gradually and softly open the autoclave enabling the steam to escape and allow the load to cool further. Hot Air Oven A hot air oven is electrically operated and should be equipped with a fan to ensure uniform temperature inside. The required temperature for sterilization is generally 160oC for one hour (Fig. 7.5).

Figure 7.5: Hot air oven

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Operating Instructions • Arrange the material to be sterilized loosely and evenly on the racks of the oven allowing free circulation of air and thereby even heating of the load. • Do not pack the load tightly since air is a poor conductor of heat. • Switch on the power supply and control the temperature of the oven by adjusting the thermostat. • Note the time when the desired temperature is reached (heatingup time). • Hold the load in the oven at this temperature for a definite period of time (holding period). This is usually 60 minutes at 160oC. • Do not overheat since it would char the cotton plugs and paper wrappings. Autoclaves and hot air ovens can be used for disinfection of infectious waste before it is discarded. In addition, waste can be disposed of by boiling in detergent or by burial. Boiling In the absence of an autoclave, most specimen containers can be boiled in water having detergents to decontaminate. This process kills the vegetative bacteria but fails to destroy the spores and certain viruses. The easiest way to get best results is to add washing powder or sodium carbonate crystals, 60 grams to one litre of water in a big container and boil specimen containers in it for a minimum of 30 minutes. Disinfection Disinfection can be undertaken either chemically or by boiling. Boiling is an effective method to disinfect equipment, e.g. needles and syringes, if autoclaving facilities are not available. Equipment which has already been cleaned should be boiled for 20 minutes. Chemical disinfection is used for heat-sensitive equipment that is damaged at high temperatures. Commonly-used chemical disinfectants include chlorine releasing compounds; ethyl and isopropyl alcohol, quaternary ammonium compounds and gluteraldehyde.

Sterilisation 33 The synopsis of a few commonly-used disinfectants is given in Table 7.1. Preferred methods of sterilization for common articles are given in Table 7.2. Decontamination of some of the commonly reusable equipment has been briefly presented in Table 7.3. Table 7.1: Disinfectants and their mode of application* Target

Disinfectant

Strength to use (disinfectant/ material V/V)

Application

Time of exposure

Skin

Ethanol Iodine Povidone iodine Quaternary ammonium comp

70% 1% 1%

Direct Direct Direct

2 minutes 2 minutes 2 minutes

Direct

2 minutes

Blood

Cresol (pH 9) 5% Ca hypochlorite 1%

2:1 2:1

6 hours 6 hours

Urine

Cresol (pH 9)

5%

1:1

4 hours

Sputum

Cresol (pH 9)

5%

1:1

4 hours

Faeces

Cresol (pH 9) Hypochlorite (Na/Ca) Ca hydroxide

5% 1%

2:1 3:1

6 hours 6 hours

20%

2:1

6 hours

Work benches

Lysol Cresol Hypochlorite Chloramine-T

5% 1% 5%

Direct Direct Direct Direct

4 4 4 4

Glassware

Hypochlorite

1%

Direct

4 hours

0.1% 70%

Direct Direct

4 hours 4 hours

Lab Hypochlorite instruments Isopropanol

hours hours hours hours

* Based upon: Basics of quality assurance: WHO/EMRO, 1992, page 162

Biohazard Waste Management Waste is defined as any solid, liquid or gaseous material that is no longer used and will either be recycled, disposed of or stored in anticipation of treatment and/or disposal.

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Practical Manual of Medical Microbiology Table 7.2: Preferred methods of sterilization for common-use articles Autoclaving

Hot air oven

Animal cages Sugar tubes Lab. coats Cotton Filters Instruments Culture media

Glass ware Beakers Beakers Petridish Pipette Slides Glass syringes Test tubes Powders

Rubber

Wood

Gloves, stopper, tubing

Tongue depressor, applicator

Glass Slides, syringes, test tubes Enamel metal trays Wire baskets Table 7.3: Disinfection of specific equipment Container/material

Method of choice for decontamination

Alternative method of decontamination

Reusable stool container

Autoclaving 121oC for 30 minutes

Fill the jar having stool with 5% solution of phenol and keep for 24hours

Empty into lavatory*

Empty into lavatory*

Autoclaving

Boiling in detergent

Reusable containers of CSF, pus, sputum

Urine bottles (after Autoclaving emptying in lavatory*)

Fill with 2% phenol or 1% bleach, leave for 4 hours, clean with detergent

Blood containers

Autoclaving

Soak overnight in strong disinfectant (5% cresol; 1% Ca hypochlorite, 1:2 V/V)

Glass microscope slides**

Autoclaving

Soak overnight in 5% phenol

* If the lavatory is connected to a septic tank, phenol or other antiseptics should not be put into the lavatory. ** Glass microscope slides which have been used for the diagnosis of tuberculosis should be discarded after keeping them soaked in detergent overnight.

Sterilisation 35 Storage Prior to disposal, all biohazardous waste should be maintained and stored separately from the general waste stream and from other hazardous wastes. The containers used to store biohazardous waste should be leak-proof, clearly labelled with a red or orange universal biohazard symbol and sealed tightly when transported. In certain cases, it may be necessary to double-bag the waste to prevent leakage. Any biohazardous sharps, such as infectious needles and scalpels, must be placed in containers that are puncture-resistant, leak-proof on all sides and the bottom, and close-able. These containers can then be placed in a standard biohazard bag. Disposal Options There are three main disposal options: Render the waste noninfectious by autoclaving and dispose it in the general waste stream. If autoclaving is not possible, decontaminate with chemical disinfectants or by boiling for 20 minutes before disposal. On-site incineration, if possible. Transportation of locally-generated waste to a distant appropriate facility. Incineration is the preferred disposal option. Not only does this method render the waste noninfectious but it also changes the form and shape of the waste. Sterilization is an effective method for decontaminating waste, but it does not alter the appearance of the waste. Steam sterilization in an autoclave at a temperature of 121oC for at least 15 minutes destroys all forms of microbial life, including high numbers of bacterial spores. This type of complete sterilization can also be accomplished using dry heat which requires a temperature of 160-170oC for 2-4 hours. However, it must be ensured that heat comes in contact with the material to be rendered sterile. Therefore, bottles containing liquid material should have loosened caps or cotton plug caps to allow for steam and heat exchange within the bottle. Biohazard bags containing waste should be tied loosely. Once sterilized, biohazardous waste should be sealed in appropriate

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containers, labelled as disinfected waste and disposed of in an approved facility. Biological waste should be clearly labelled prior to disposal and complete records should be maintained. Burial It is not a recommended decontaminating process. However, it does prevent the infectious material from becoming a reservoir of infection if properly buried. It requires digging a pit of almost 5 meters depth and 2 meters width and having a tightly fitted heavy lid on top. Disposable containers with clinical material are thrown daily into it and the lid is replaced immediately after throwing the specimens. Once a week, the refuse is covered with a layer of quicklime. If quicklime is not available, the refuse is covered with almost 10 cm thick layer of dried leaves once a week.

8 Hanging Drop Preparation ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION This experiment is mainly used to study the motility of bacteria. It is also useful to study the shape, relative size and arrangement of bacteria, protozoan parasites, sperm motility and stool examination. The term hanging drop is self explanatory, after the preparation the drop hangs from the coverslip. The hanging drop provides enough space for the movement of Bacteria Unlike wet mount where a thin film is available for movement. Requirements 1. Young broth culture 2. Clean cavity slide (depression slide). cavity slide is slide with central depression or concavity (Figs 8.1 and 8.2). 3. Clean cover slip 4. Vaseline 5. Spirit lamp 6. Wire loop 7. Compound microscope.

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Figure 8.1: Cavity slides

Figure 8.2: Cavity slide with hanging drop

Procedure (Fig. 8.3) • Take a clean cover slip and apply a little of Vaseline to the corners. • Place a drop of broth culture on the centre of the cover slip using a sterile wire loop. • Invert a clean cavity slide gently over the cover slip, the concavity facing the drop (i.e. downwards) See that the drop does not touch the slide. • Quickly and carefully, turn the slide over. So that the coverslip is upper most and the drop is suspending from the coverslip. • Lower the substage condenser to reduce the light.  Partially close the iris diaphragm if necessary.

Hanging Drop Preparation 39

Figure 8.3: Hanging drop preparation

• Focus the drop under low power objective so that the edge of the drop is exactly in the centre of the microscopic field. • Turn to high power objective and focus the edge of the drop. • Study the motility, shape, relative size and arrangement of the organisms. Observation (Fig. 8.4) Motile and non motile organisms may be observed.

Figure 8.4: Edge of the drop with motile bacilli

Notes The edge of the drop is focussed because of the following reasons. 1. The contrast or optical aberration due to the different refractive indices (Water and glass) is less at the edge of the drop. 2. The organisms move in a horizontal direction at the edge of the drop, contrary to the vertical movement seen at the centre of the drop.

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3. Oxygen tension is more towards the edge of the drop and hence aerobes tend to accumulate at the edge of the drop. Bacterial Motility Bacteria exhibits true motility and false motility. When a bacterium changes its position and moves in different directions, we can consider it as true motivity. False mobility is mainly due to Brownian movement, i.e. the organisms exhibit an oscillatory movement without change of position, due to the bombardment of water molecules True motility is different types. 1. Darting (missile like), e.g. Vibrio, Pseudomonas 2. Tumbling motility, e.g. Salmonella, Esch.coli. Examples of Motile Bacilli • • • •

Salmonella Vibrio cholerae Pseudomonas. Proteus.

Examples of Non-motile Bacilli • • • •

Klebsiella Shigella Clostridium welchii Haemophilus influenzae.

Motility of Anaerobic Bacteria Motility of anaerobic bacteria can be checked by inoculating them into a semisolid medium. Motile anaerobes are seen to spread at the deep layers of the medium where oxygen content is less or neglibile. Another microscopic method used for anaerobic motility observation is closed Capillary tube method. Liquid growth from a liquid anaerobic culture medium (Thioglycollate broth) is taken in a Capillary tube, close the ends of cappillary tube by heating, observe the capillary tube under microscope.

9 Preparation and Fixation of Smears ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

A proper fixed smear is essential for many staining procedures like Gram’s staining and Acid fast staining. A good smear will provide reliable information if they are stained systematically, labelled properly, spread evenly and fixed with care.

Note • It is recommended to prepare only one smear on a single slide. • Slides with positive AFB Smears should always be discarded and never reused. • Precaution should be taken when handling infectious material. • The Smears should be spread evenly covering an area of about 15-20 mm diameter on a slide. Requirements 1. 2. 3. 4. 5. 6. 7.

Clean, clear, grease free glass slide Wire loop Burner or spirit lamp (Fig. 9.1) Sterile normal saline or distilled water Glass marking pencil 70% methanol or Ethanol (for Alcohol fixation of M. tuberculosis) Absolute alcohol (for alcohol fixation other Bacteria)

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Figure 9.1: Spirit lamp

Procedure Step 1—Making of smears Step 2—Drying of smears Step 3—Fixation of smears Step 4—Labelling of slides Making of Smears Smears can be made out of various clinical specimens and bacterial cultures. 1. Bacterial cultures: Smear can be prepared by the following procedure. • Sterilise the wire loop by showing it in the flame, make it red hot and allow to cool. • Place one or two drops sterile saline on a clean slide. • Using the sterile wire loop pick up a small portion of the isolated Bacterial colony.

Preparation and Fixation of Smears 43

2. 3.

4. 5. 6.

• Emulsify the colony in the saline and spread evenly on the slide in a circular manner. Pus: Spread the purulent material thinly using a sterile wire loop. The flame sterilized loop must be allowed to cool before it is used. Sputum: Use a piece of clean stick to transfer and spread purulent and caseous material on a slide. Dip the stick in a disinfectant solution before discarding. Swab: Roll the swab on a slide. Rolling the swab prevents damage to the pus cells. Faeces: Use a piece of clean stick to transfer pus and mucus to a slide. Decontaminate the stick before discarding. CSF: Make on evenly spread smear of a drop of purulent CSF or the sediment from a centrifuged sample on a slide using a sterile wire loop. (AFB is difficult to detect in CSF. The chances of finding Bacteria are increased if the CSF it centrifuged for 20-30 min and several drops of sediment are used).

Drying of Smears After making smears, the slides should be kept in a safe place to air dry. Protect the slide from flies and dust. If the smears can not be stained immediately, they should be Fixed and stored in a covered box. Fixation of Smears The purpose of fixation is to preserve microorganisms, and to prevent smears being washed from slides during staining. Smears are fixed by heat, alcohol or by other chemicals. Heat fixation: The air dried smear is fixed by passing the slide (Smeared slide up) rapidly over the burner by holding the slide with the thumb and index finger (Don’t over heat). The slide is them allowed to cool (Fig. 9.2). The aim of this fixing is to coagulate the aluminous material whereby the film adheres better to the glass slide and thus may not be detached in the subsequent staining process. Heat fixation is commonly used for Gram’s staining.

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Figure 9.2: Heat fixed smear

Note: After passing the slide through the flame three times, it should be possible to touch the slide on the back of your hand without the hand feeling uncomfortably hot. The disadvantages of heat fixation are : 1. It damages leucocytes and 2. M. tuberculosis is not killed by this method. Alcohol fixation: This fixation is more bactericidal than heat fixation, it kills M. tuberculosis. Another advantage of alcohol fixation is, it does not produce damage to the puscells, and they are well preserved. Alcohol fixation is recommended for fixing smears containing instracellular Gonococcus and Meningococcus. method of alcohol fixing smear is as follows. 1. Allow the smear to air-dry completely. 2. Keep the slide on a staining rack. 3. Add one or two drops of alcohol on the smear. 4. Leave the alcohol on the smear for 2 minutes to dry the alcohol on the smear. Example other chemical fixations: • 40 gm/l potassium permagnate is recommended for fixing smears containing Anthrax bacilli. • Formaldehyde Vapour. For smear containing Mycobacterium. Labeling of Slides Using a grease pencil mark the area of smear. This will be useful while focusing under microscope. Label the slide with date, name and numbers.

10 Methylene Blue Staining ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION This is a simple staining procedure, both background and object will stain blue. Requirements Methylene blue 0.3 gm Distilled water 100 ml Dissolve the dye in water. Filter through a filter paper. Staining Method Make a smear on a glass slide, dry in air and fix by passing it over the flame of a burner 3-4 times. Stain for one minute by pouring methylene blue solution over the smear. Wash with water, blot dry and examine under the oil immersion of light microscope (Fig. 10.1).

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Figure 10.1: Methylene blue staining of yeast cells

Uses The stain is used to make out clearly the morphology of the organisms, e.g. Yersinia pestis in exudate, Haemophilus influenzae in CSF and Gonococci in urethral pus.

11 Indian Ink Staining ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION This is a negative staining, useful for demonstration of capsule of bacteria and yeasts. The capsule will not take up the stain and the remaining areas of the field will be darkly stained. Staining Procedure • Place a loopful of India ink on the side of a clean slide. • A small portion of the solid culture is suspended in saline on the slide near the ink and then emulsified in the drop of ink, or else, mix a loopful of liquid culture of specimens like CSF with the ink. • Place a clean cover slip over the preparation avoiding air bubbles. • Press down, or blot gently with a filter paper strip to get a thin, even film. • Examine under dry objectives followed by oil immersion. Observation To demonstrate the capsule which is seen as an unstained halo around the organisms distributed in a black background. This is

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employed for diagnosis especially for Cryptococcus neoformans and Pneumococcus (Fig. 11.1).

Figure 11.1: Indian ink preparation of Pneumococcus

12 Gram's Stain ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION In 1884 Hans Christian Gram described this method of staining, which is the most important stain in routine bacteriology. It is differential stain used for the identification of Bacteria. Requirement a. Crystal violet b. Gram’s iodine

c . Absolute alcohol/acetone d. Dilute carbol Fuchsin

Composition: Crystal violet and distilled water. Act as mordant. Composition – Iodine, potassium iodide and distilled water Decolouriser. Counter stain. Carbol fuchsin and distilled water.

Procedure (Fig. 12.1) 1. Preparation of film – A thin uniform smear is absolutely essential. 2. Fixation of the smear – The smear is dried in air and fixed by passing the slide (Smeared slide up) rapidly over the spirit lamp

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Figure 12.1: Gram’s stained smear with GPC, GPB, GNC, and GNB (For colour version, see Plate 1)

3. 4.

5.

6. 7. 8. 9.

or Bunsen burner by holding the slide with the thumb and index finger. (DON’T OVER HEAT). The slide is then allowed to cool. The object of this fixing is to coagulate the albuminous material whereby the film adheres better to the glass and thus may not be detached in the subsequent processes. A film should never be heated over not be detached in the subsequent processes. A film should never be heated over the flame until it is thoroughly dried up. Cover the smear with crystal violet solution and allow to remain on the slide for 1 minute. Discard the cyrstal violet stain and hold the slide at a steep and wash off the residual stain with an excess of iodine solution. Cover the smear with fresh iodine solution and leave it for 1 minute. Rinse the smear with absolute alcohol and continue application until no more colour appears to flow from the preparation (about 15-20 seconds).if acetone is used for decolorisation, wash immediately. Wash with water. Cover the smear with dilute carbol fuchsin and allow it to remain for 1 minute. Rinse with water and dry in air or between blotting paper. Then examine under oil immersion.

Gram's Stain 51 Observation (Fig. 12.2) • Gram positive organism stains violet. • Gram negative organism stains pink.

Figure 12.2: Gram’s staining steps (For colour version, see Plate 1)

COCCI

Gram positive cocci

Gram negative cocci

- Staphylococcus arranged in clusters - Sterptococcus in chains. - Pneumococci is lanceolate, diplococci and Capsulated. - Neisseria-bean shaped diplococci.

BACILLI Gram positive bacilli: Corynebacterium diphtheriae, Clostridium spp. Gram negative bacilli: Salmonella, Shigella, Proteus and Pseudomonas. Comments Gram stain divides the bacteria into two catergories, depending upon whether they can be decolorised with alcohol after staining with

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crystal violet and iodine. Those that resist decolourisation remain violet in colour and are designated as Gram positive, and those that are decolourised and take up the counter stain such as dilute carbol fuchsin appear pink and are termed as Gram negative. The suggested reasons for this differentiation into two categories are: 1. Gram positive organisms have a more acidic protoplasm (pH 2.0) than the Gram negative organisms (pH 5.0), there by having greater affinity for basic dyes. 2. Cell wall theory—After staining with crystal violet and iodine, a dye iodine complex is formed within the cell. This complex is insoluble in water but moderately soluble and dissolvable in alcohol. Under the action of decoloriser the dye and iodine diffuse freely out in Gram negative cell but not from Gram positive cell, presumably because the latter surface is less permiable to the decoloriser or its iodine solute. 3. Cyto plasmic theory—Gram positivity depends upon the integrity of cellular structure and the presence in the cell of a specific magnesium ribonucleate complex. Thus Gram positive bacteria become Gram negative if they are ruptured mechanically or if this ribonucleate is removed.

13 Ziehl-Neelsen’s Stain (Acid Fast Stain) ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Mycobacteria have the power of retaining certain stain even when they are decolorised by mineral acids. This is known as “Acid fastness”. This acid fastness appears to be due of presence of lipid (Mycolic acid) and integrity of the cell wall. The improvement of acid fast staining was done by Ziehl and Neelsen. It is a differential stain used to differentiate bacteria into acid fast and non-acid fast. Requirements a. Strong carbol fuchsin-composition: Basic fuchsin, absolute alcohol, phenol and water. b. Sulphuric acid 20% solution. c. Absolute alcohol (Used only if the genitourinary specimens are processed) d. Methylene blue-composition: Methylene blue in alcohol and potassium hydroxide and water. Procedure 1. Preparation of smear: A portion of the mucopurulent material is taken on a slide and a smear is made and dried.

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2. “Fixation of the smear”-fix the smear by gently passing the slide over the flame. 3. Flood the slide with strong carbol fuchsin and heat until steam rises. Allow the preparation to stain for 5 to 7 minutes. Heat being applied at intervals until steam rises and stops it and continue in this way for 5-7 minutes (never heat too much to produce boiling and charring. The stain must not be allowed to evaporate and dry on the slide, if necessary, pour more carbol fuchsin on the slide. 4. Wash in running tap water. 5. Decolourise the smear with sulphuric acid (20%) by dipping the slide in the acid solution in a wide mouth jar. Decolourisation should be continued till the film becomes faintly pink. Decolourisation must be done in stages and generally requires about 10 minutes. 6. Wash in running tap water. 7. Counter stain the smear with methylene blue for 1-2 minutes. 8. Wash in running tap water. 9. Blot dry and examine with the oil immersion objective. Observation (Fig. 13.1) • Acid fast bacilli – Pink • Tissue and other organisms – Blue • Mycobacteria are acid fast bacilli and most of the other bacteria are non-acid fast. • Nocardia is another bacterium which shows acid fastness. Notes Acid fast bacteria are very difficult to stain. Hence heating is necessary while staining which act as a mordant and makes the tough lipid cell wall of the bacterium permeable to the dye. In cases of genitourinary specimens the absolute alcohol or 3% acid alcohol can be used as decolouriser. In case of suspected lepra bacilli, only 5% sulphuric acid is to be used as decolouriser because it is less acid fast. Most important acid fast bacteria of medical importance are Tubercle bacilli (Human, bovine, atypical) lepra bacilli and smegma bacilli, M. smegmatis is not pathogenic to human beings. Tubercle

Ziehl-Neelsen’s Stain (Acid Fast Stain) 55

Figure 13.1: Z-N stained sputum smear with AFB (pink) (For colour version, see Plate 2)

bacilli are both acid and alcohol fast, Smegma bacilli is acid fast but not alcohol fast. Lepra bacilli are acid fast to a lesser degree and alcohol fast. Some of the suspected tuberculosis cases may fail to show acid fast bacteria by ordinary Z- N stain methods. In such cases other methods like concentration techniques and direct flourescent staining may be helpful. In concentration technique the specimen is digested suitably with alkali at neutral pH and the digested material is centrifuged. The smear made from the deposit are stained by Z-N-stain and observed.

14 Albert’s Stain ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Some bacteria possess granules in the cytoplasm these granules are known by different names like metachromatic granules or volutin granules or Babes-Ernst granules or polar bodies. Demonstration of these granules is useful in identification of the Bacterial Species. These Granules are made up of glycogen, Starch, lipids and polymetaphosphates. The granules consisting of polymetaphosphate are called voluntin granules or Babes-Ernst granules. Babes-Ernst granules are predominant in Corynebacterium diphtheriae. For demonstration of the volutin granules three special staining methods are available they are Alberts staining, Neisser’s staining and Ponder’s staining. Albert’s staining is the common staining procedure in the routine microbiology laboratories. This staining method is useful for the identification of Corynebacterium diphtheriae. The granules take up a different staining colour from that of the stain itself and hence the name metachromatic granules. Usually each bacillus contains 2 to 6 such granules, mainly in the poles of the bacilli. These are reserve food materials and appear more prominent during starvation.

Albert's Stain 57 Requirement 1. Albert’sStain Toludine blue - 1.5 gm Malachite green - 2 gm Glacial acetic acid - 10 ml Ethyl alcohol (95%) - 20 ml Distilled water - 1litre Dissolve the dyes in the alcohol and add to the water and acetic acid. Allow to stand for one day and then filter. 2. Albert’sIodine Iodine - 6 gm Potassium Iodide - 9 gm Distilled water - 900 ml Method • Prepare a smear, allow to dry and fix with gentle heat. • Cover the smear with Albert’s stain and allow to act for 3-5 minutes. • Discard Albert’s stain and hold the slide at a steep slope and wash off the residual stains with an excess of Albert’s Iodine Solution Cover the Smear with fresh Albert’s iodine solution and allow to act for 1 minute. • Wash with water, blot dry. • Observe under oil immersion objective. Observation (Fig. 14.1) • • • •

Granules bluish black, the protoplasm green Granules mainly seen in the poles. Other organisms usually pale green. The diptheria bacilli show typical arrangement of letters like L,V,N or like Chinese letters.

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Figure 14.1: Albert stained smear of corynebacterium (For colour version, see Plate 2)

15 Leishman’s Stain ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION This stain is a modification of Romanowsky stain. It is very useful for the demonstration of protozoa in blood films like Malaria parasite. Requirements 1. Leishman staining solution 2. Distilled water or buffer solution Leishman’s Stain is prepared by taking 0.15 gm of Leishman’s powder and dissolving the powder in 100 ml methanol. Distilled water should be neither acid nor alkaline. Any slight variations from neutrality may alter considerably the colour of granules in white blood corpuscles and give rise to supposed pathological appearances in cells which are really normal. This can be over come by using a buffer solution. Method • Make thin film of blood smear. • Pour the undiluted stain on to unfixed film and allow it to act for 1 min.

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Figure 15.1: Leishman stained blood smear (For colour version, see Plate 3)

• Using a pipette add double the volume of distilled water/buffer to the slide mixing the fluids by alternately sucking them up in the pipette and expelling them. Allow the diluted stain to react for 12 min. • Flood the slide gently with distilled water. • Remove the excess water and blot dry. Note Methyl alcohol present in the stain acts as a fixative and fixes the film to the slide.

Leishman’s Stain 61 Observation (Fig. 15.1) RBC : Red WBC : Blue Malarial parasite : Inside RBC with blue cytoplasm and ruby red nucleus.

16 Preparation and Cleaning of Glassware ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

NEW GLASSWARES New glass wares may contain resistant spores which may be present in packing material and new glass wares tend to give off free alkali which may be sufficient to interfere with the growth in certain organisms. To overcome these problems the following cleaning methods should be followed (Figs 16.1 to 16.4). • Place in 1% HCl overnight • Wash with tap water • Wash in distilled or deionised water • Autoclave. Used Glasswares • Reusable glass ware should be autoclaved before cleaning. • The discarded cultures and their containers are then placed in a hot detergent solution clean with a suitable brush. • Rinse with deionized water • Drain and dried in a hot air oven • Dry sterilise at 160 oC for 3 hours.

Preparation and Cleaning of Glassware 63

Figure 16.1: Laboratory wares (plastic/glass)

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Figure 16.2: Laboratory wares (plastic/glass)

Chromic Acid Cleaning This cleaning of glassware is mainly for biochemical work. • Remove any grease with petroleum. Wash with warm tap water. • Place in dichromate Sulphuric acid cleaning solution for 12-24 hours. • Remove and wash by density in hot tap water at least four times and in distilled water twice.

Preparation and Cleaning of Glassware 65

Figure 16.3: Preparation of glass Pasteur pipette

Figure 16.4: Plastic Pasteur pipettes

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• Dry in oven if the glass ware is not used for accurate Volumetric purposes. Cleaning of Pipettes • If contaminated with infected material, place the used pipette into disinfectant solution and leave until convenient to wash. • Rinse in tap water. • If necessary, keep overnight in detergent or dichromate–Sulphuric cleaning fluid. • Wash with tap water followed by deionized water. • The top end of the pipette is plugged with cotton-wool. Press it entirely with in the end of the pipette so that there are no protruding strands of cotton to prevent close fitting of a rubber teat or other pipette filling device which may later be attached to operate the pipette. • To sterlise the pipettes, Pack them in aluminium or copper cylinders. Place in a hot air oven at 160oC for 3 hours. Note: Accurately caliberated volumetric glassware should never be heated in an oven, since the expansion and contraction of the glass makes the graduations inaccurate.

17 pH in Microbiology ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Microorganisms are sensitive to the varying pH of the external environment. Where there is an optimum pH for the growth and multiplication the microorganisms will survive and flourish. So when micro organisms are cultivated in the laboratory it should be noted that the medium should have an optimum pH. Media should be adjusted as far as possible to the pH optimal for the growth of the organism concerned. Most pathogenic bacteria have a fairly restricted PH range and grow best around pH 7.3, i.e. at a slightly alkaline reaction. Methods Used for Measurement of pH a. The pH meter: The accurate method of measuring pH is with a pH meter. It is easy and quick to use. Care must be taken in its maintenance. b. pH indicator dyes: Indicator dyes are substances that will change in colour with variations in the pH of the solution in which they are dissolved. Examples of indicative dyes and pH range are given in the Table 17.1.

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Practical Manual of Medical Microbiology Table 17.1: Indicative dyes and pH range Indicator Thymol blue Bromophenol blue Bromocresol green Methyl red Litmus Bromocresol purple Bromothymol blue Neutral red Phenol red Cresol red Thymol blue Phenol phthalein Thymol phthalein

Range of pH 1.2-2.8 2.8-4.6 3.6-5.2 4.4-6.2 4.5-8.3 5.2-6.8 6.0-7.6 6.8-8.0 6.8-8.4 7.2- 88 8.0-9.6 8.3-10.0 9.3-10.5

Colour change Red to yellow Yellow to violet Yellow to blue Red to yellow Red to yellow Yellow to violet Yellow to blue Red to yellow Yellow to purple pink Yellow to violet red Yellow to blue Colourless to red Colourless to blue

pH Indicator Papers The simplest method of determining the pH of a solution is use of commercially available pH indicator papers. These papers are impregnated with an indicator that gives a change of colour over a specific or general range of pH. The Paper can simply be dipped in the solution to be tested or a drop the solution can be withdrawn by a wire crop or pasteur pipette are placed on the paper. The resulting colour is compared with the chart supplied with the papers. Comparater and Capillator Methods These methods are not available now so not used.

18 Bacteriological Media ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

The role of suitable quality culture media for cultivation of microorganisms cannot be over emphasized. The success of isolation of aetiological agents depends on the quality of the medium. Only in exceptional cases, can an organism be identified on the basis of its morphological characteristics alone. Types of Media Bacteriological media can be broadly sub-divided into four categories. 1. Ordinary Culture Media These are routinely employed in a laboratory, e.g. nutrient broth, nutrient agar, infusion broth and lysate media. 2. Enriched Media Certain organisms do not grow on ordinary nutrient media. They require growth-promoting ingredients such as blood, glucose, serum, egg, etc. The media containing ingredients which enhance their

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growth-promoting qualities are enriched media, e.g. blood agar, chocolate agar and Loeffler medium. 3. Enrichment Media Enrichment media are liquid media containing chemical constituents which inhibit some normal flora and allow pathogens which may be present in very small number in the specimen, to grow unhampered and thus enriching them. Isolated colonies of these organisms may be obtained by subculturing onto solid media. An example of enrichment media is selenite broth used for primary isolation of enteric bacteria. 4. Differential and Selective Media Differential media have got some chemical constituents which characterize different bacteria by their special colonial appearances in the culture, e.g. MacConkey agar contains lactose as a substrate and neutral red as an indicator. Bacteria fermenting lactose produce acid and this will change the colour of the indicator and thus the colonies will turn red. The red lactose fermenting colonies can be differentiated from the pale non-lactose fermenting colonies. Selective media will selectively permit the growth of pathogens and inhibit the commensals. In addition, it may differentiate the pathogen from commensals that grow by the colour and opacity of the colonies, e.g. blood tellurite medium for C. diphtheriae. In addition, transport media are also frequently used to sustain the viability of organisms when a clinical specimen is to be transported from the periphery to laboratory. The transport medium prevents the outgrowth of contaminants during transit and sustains the pathogen. Cary and Blair and Stuart media are two examples of this group of media. Preparation of Media and Checking of pH Presently, a wide range of culture media are available commercially in the form of dehydrated media. These media are simply reconstituted by weighing the required quantities and by adding distilled water, as per the manufacturer’s instructions.

Bacteriological Media 71 The pH determination can be conveniently done with the use of pH indicator papers. Adjust the pH using NaOH (1 Normal) and HCl (1 Normal) Check the pH of the medium once again before use. The quantity of agar given in the formulae of media may have to be changed depending upon the quality of agar used. The concentration varies from batch to batch and should be such that will produce a sufficiently firm surface on solidification. This can be tested by streaking with inoculating wire. In some laboratories media are prepared by individual measurement of ingredients and then mixing the same (Figs 18.1 to 18.4). Nutrient Broth (Fig. 18.1) • • • •

Meat extract 10.0 gm Peptone 10.0 gm Sodium chloride 5.0 gm Distilled water 1000 ml Mix the ingredients and dissolve them by heating in a steamer. When cool, adjust the pH to 7.5-7.6. Nutrient Agar (Fig. 18.2) To the ingredients as in nutrient broth, add 15 gm agar per litre. Dissolve the agar in nutrient broth and sterilize by autoclaving at 121oC for 15 minutes. Prepare plates and slopes as required. Glucose Broth • Nutrient broth 900 ml • Glucose (10% solution) 100 ml • Dissolve 9 gm glucose in distilled water and sterilize by tyndallisation. • Add l00 ml of the glucose solution to 900 ml of sterile nutrient broth. • Dispense 60 ml each in 100 ml pre-sterilized culture bottles. • Sterilize by open steaming at l00oC for one hour.

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Figure 18.1: Nutrient broth (For colour version, see Plate 3)

Figure 18.2: Nutrient agar with bacterial colonies (For colour version, see Plate 4)

Bacteriological Media 73

Figure 18.3: Blood agar (For colour version, see Plate 4)

Figure 18.4: Colony morphology

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Blood Agar (Fig. 18.3) Nutrient agar 100 ml Sheep blood (defibrinated) 10 ml Melt the sterile nutrient agar by steaming, cool to 45oC. Add required amount of sheep blood aseptically with constant shaking. Mix the blood with molten nutrient agar thoroughly but gently, avoiding froth formation. Immediately pour into petri dishes or test tubes and allow to set. Chocolate Agar The ingredients are essentially the same as in blood agar. Melt the sterile nutrient agar by steaming and cool to about 75oC. Add blood to the molten nutrient agar and allow to remain at 75oC after gently mixing till it is chocolate brown in colour. Pour in Petri dishes or test tubes for slopes as desired. XLD Agar Xylose 3.5 gm 1–lysine 5.0 gm Lactose 7.5 gm Sucrose 7.5 gm Sodium chloride 5.0 gm Yeast extract 3.0 gm Sodium desoxycholate 2.5 gm Sodium thiosulphate 6.8 gm Ferric ammonium citrate 0.8 gm Phenol red 0.08 gm Agar agar 15.0 gm Water 1000 ml Weigh the ingredients into a flask and add distilled water. Mix the contents well and steam it for 15 minutes (do not autoclave). Cool to 56oC and pour in plates.

Bacteriological Media 75 Buffered Glycerol Saline Glycerol 300 ml Sodium chloride 4.2 gm Disodium hydrogen phosphate 10.0 gm Na2 HPO4 Anhydrous 15.0 gm Phenol red aqueous solution 0.02 per cent 15.0 ml Water 700 ml Dissolve NaCl in water and add glycerol. Add disodium hydrogen phosphate to dissolve. Add phenol red and adjust pH to 8.4. Distribute 6 ml in universal containers (screw -capped bottles of 30 ml capacity). Autoclave at 115oC for 15 minutes. Loeffler Serum Medium Nutrient broth 100 ml Serum (sheep or horse or ox) 300 ml Glucose 1.0 gm Dissolve glucose in nutrient broth and sterilize at 121oC for 15 minutes. Add serum aseptically. Mix thoroughly but gently, avoiding froth formation. Distribute in sterile test tubes or quarter ounce screw-cap bottles. Inspissate the medium in a slanting position in a water inspissator at 82oC for two hours. In the absence of an inspissator, the medium may be coagulated by standing over the top of a steam sterilizer for 6-7 minutes. Blood Tellurite Agar Agar base Meat extract 5.0 gm Peptone 10.0 gm Sodium chloride 5.0 gm Agar 25.0 gm Water 1000 ml Dissolve the ingredients and adjust the pH to 7.6. Distribute in 100 ml quantities in a bottle and autoclave at 121oC for 15 minutes.

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Glycerolated blood tellurite mixture Sterile defibrinated sheep blood 14 ml Sterile glycerol 6 ml Sterile potassium tellurite solution (1% in water) 4 ml Sterilise the glycerol in hot air oven at 160oC for 60 minutes and the tellurite solution by autoclaving at 115oC for 20 minutes. Mix the ingredients in a sterile flask, incubate for 1-2 hours. at 37oC, then refrigerate. Haemolysis is complete after 24 hours. The mixture keeps well in a refrigerator. One per cent solution of good quality tellurite is sufficient but 2% of some batches may be required. Preparation of complete medium Glycerolated blood tellurite mixture 24 ml Agar base 100 ml Melt the agar, cool to 45oC, add blood and tellurite and pour in sterile petri dishes.

19 Inoculation of Culture Media ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

When inoculating or seeding culture media an aseptic (sterile) technique must be used to: • To prevent contamination of cultures and specimen. • To prevent infection of the laboratory worker and the environment. Aseptic Technique (Fig. 19.1) • Flame sterilize wire loops, straight wires, and metal forceps before and after use whenever possible, use a hooded Bunsen burner (Fig. 19.2). Note: To prevent the release of aerosols, wire loops should be well made. Aerosols can also be released when spreading inoculation media containing air bubbles. • Flame the necks of specimen bottles, culture bottles, and tubes after removing and before replacing caps, bungs, or plugs. • When inoculating, do not let the tops or caps of bottles and tubes touch an unsterile surface. This can be avoided by holding the top or cap in the hand. • Always use racks to hold tubes and bottles containing specimens or culture media.

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Figure 19.1: Aseptic inoculation of culture medium

Figure 19.2: Bunsen burner

Inoculation of Culture Media 79 • Make slide preparations from specimens after inoculating the culture media. • Decontaminate the work bench before starting the day’s work and after finishing. • Use a safety cabinet when working with hazardous pathogens. • Wear protective clothing, wash the hands after handling infected material, and never mouth-pipette, eat, drink, or smoke in the laboratory. Making a Wire Loop Loops must be made correctly to ensure inocula are well spread, and to prevent the release of aerosols from long and springy loops or loops that are not completely closed. The length of wire from the loop to the loop holder should be about 50 mm. The method of making a wire loop is as follows (Fig. 19.3):

Figure 19.3: Preparation of wire loop

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1. Cut a piece of wire about 125 mm in length and thickness (swg) 26 or 27, Wind it around a loop holder. 2. Using a pair of scissors, cut off one arm of the wire leaving the loop and about 50 mm of wire. Bend the loop back to make it central using a pair of forceps. 3. Insert the wire in a loop holder. Make sure the loop is completely closed. Note: When sterilising a wire loop, hold it in the blue part of a Bunsen burner flame allow the loop to cool before using it. Inoculation of Media in Petri Dishes (Fig. 19.5) The technique used to inoculate media in petri dishes (plates) must provide single colonies for identification and to see whether a culture is pure or mixed, i.e. consisting of a single type of organism or several different organisms. A pathogen must be isolated in pure culture before it can be identified and antimicrobial sensitivity tested. The inoculation of media in petri dishes is referred to as ‘plating out’ or ‘looping out’. It is not necessary to use whole plates of media. Considerable savings can be made by using a half or even a third of a plate (especially if the medium is a selective one). The area of medium used must be sufficient to give separate colonies (Fig. 19.4). Before inoculating a plate of culture medium, the surface must be dried first otherwise single colonies will not be formed. Usually 3040 minutes incubation at 37oC is adequate.

Figure 19.4: Incubator (For colour version, see Plate 5)

Inoculation of Culture Media 81

Figure 19.5: Inoculation of media

To inoculate a plate, apply the inoculum to a small area of the plate (‘the well’) using a sterile wire loop or swab of the specimen held with sterile forceps. Flame sterilize the loop. When cool (or using a second sterile loop), spread and thin out the inoculum. This will ensure single colony growth. Inoculation of Slopes To inoculate slopes such as Dorset egg medium or Loeffler serum, use a sterile straight wire to streak the inoculum down the centre of the slope and then spread the inoculum in a zig-zag pattern. To inoculate a slope and butt medium, such as Kigler iron agar, use a sterile straight wire to stab into the butt first and then use the same wire to streak the slope in a zig-zag pattern.

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Inoculation of Stab Media (deeps) Use a Sterile straight wire to inoculate a stab medium, for example mannitol motility medium. Stab through the centre of the medium taking care to withdraw the wire along the line of inoculum without making further stab lines. Inoculation of Fluid Media Broths and other fluid media are inoculated using a sterile wire loop, straight wire, or Pasteur pipette depending on whether the inoculum is colonial growth or a fluid culture or specimen. A straight wire is used to inoculate Koser’s citrate broth to prevent any carry over of medium. If using a wire loop to subculture colonies, hold the bottle or tube at an angle and rub the loop against the side of the container below the level of the fluid. Labelling of Inoculated Media Using a grease pencil or marker pen, label inoculated media with the date and the patient’s number. Always label the base of a culture plate. A slope should be labelled on the underside of the media so that the wording does not obscure the culture. A stab culture should be labelled above the level of the agar. If a plate is to be incubated anaerobically it should be marked ‘An’ or if in a carbon dioxide atmosphere it should be marked ‘CO2’. Incubation of Cultures Inoculated media should be incubated as soon as possible. A delay in incubation can effect the viability of pathogens especially anaerobes, pneumococci, meningococci, gonococci, and Haemophilus influenzae. It can also increase the risk of plates becoming contaminated from small insects and dust especially in the dry season. Uninoculated and inoculated media must be protected from sunlight. Microorganisms require incubation at the temperature and in the humidity and gaseous atmosphere most suited to their

Inoculation of Culture Media 83 metabolism. The length of time of incubation depends on how long an organism takes to develop the cultural characteristics by which it is recognized (Fig. 19.5). Temperature of Incubation The temperature at which a microorganism grows best is referred to as its optimum temperature. The temperature below which growth stops (not necessarily resulting in death) is called the minimum temperature, and that above which growth stops and death occurs is called the maximum temperature. Note: The thermal death point of microorganism is the lowest temperature above the optimum at which death will occur in a given time. It is influenced among other factors by the culture medium, age of culture, and moisture. The temperature selected for routine culturing is 35-37oC with most microbiologists recommending 35oC in preference to 36oC or 37oC. In general, the growth of microorganisms is more affected by slight rises above their optimum temperature than by reductionsbelow it.

20 Anaerobic Cultivation ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Depending on oxygen requirement, miroorganisms can be classified into 4 groups: 1. OBLIGATE AEROBES Those bacteria which require free oxygen for their growth. Ex.: a. Alkaligenes faecalis. b. Moraxella. c . Brucella 2. OBLIGATE ANAEROBES Those that can live and grow only in the absence of oxygen, i.e. under high reducing intensity and low redox potentials Ex.: a. Treponema denticola b. Clostridium tetani c . Clostridium botulinum. 3. FACULTATIVE ANAEROBES These bacteria which can live and grow in the presence as well as in the absence of oxygen. Most of the bacteria come under this group.

Anaerobic Cultivation 85 Ex.: a. Staphylococci. b. Esch.coli. 4. MICROAEROPHILIC ORGANISMS Those that grow just in the presence of trace of oxygen. Ex.: a. Vibrio sputorum b. Campylobacter. SOURCE OF ANAEROBES Anaerobes occur in a variety of human body sites. They are usually present as normal commensals without causing any harmful effects. But under predisposing factors, i.e. whenever there is a low-redox potential, they can act as pathogens, thus producing endogenous infections. During infection, they are isolated from various specimens like, Faeces, Blood, Pus, Exudates, etc. Methods of Creating Anaerobiosis . I By using certain gases, like, a. 100% H2 b. 90% H2 + 10% CO2 c . 80% H2 + 10% N2 + 10% CO2 II. Addition of reducing agents to the media, like, a. Thioglycollate (Fig. 20.1) b. 10%. Ascorbic acid. c . Meat particles as in Robertson’s cooked medium. d. Palladium Chloride. e . Cysteine, etc. III. Growing anaerobes deep in agar butts. IV. Simultaneous growth of aerobe and an anaerobe. V. Biological method by employing germinating seeds. VI. A Simple method, devised by Buchner employs a filter paper having the same diameter as a petridish. This filter paper is placed on the top of one half of the petridish and a mixture of pyrogallic acid, and Sodium hydroxide in dry powder form is spread on it. Then the other half of the petridish containing

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Figure 20.1: Thyoglycollate medium (For colour version, see Plate 5)

the media with inoculated specimen is inverted over the filter paper and the edge is sealed tight with molten wax. The alkaline-pyrogallate mixture will absorb all the O2 present within the closed system, thus creating complete anaerobiosis. VII. Utilization of Anaerobic Jars (Fig. 20.2). Ex.: 1. McIntosh and Filde’s Jar 2. Gaspak jar

Figure 20.2: Candle jar (left) anaerobic jar (right)

Anaerobic Cultivation 87 McIntosh and Filde's Anaerobic Jar This jar is used to provide complete anaerobic environment for the cultivation of anaerobic bacteria like clostridia, etc. It is a stout metallic or glass jar with a metal lid. Inoculated culture plates are placed inside the jar and the lid is clamped tight. The lid has 2 tubes: (1) Outlet tube which is connected to a vacuum pump to evacuate the air inside, (2) Gas inlet: which is then connected to a hydrogen supply. The lid also has 2 electric terminals, leading from which and suspended on the underside of the lid is a small porcilian spool, around which is wrapped a layer of palladinised asbestos. After filling the jar with H2, the electric terminals are connected to a current supply to heat the palladinished asbestos. This acts as a catalyst for the combination of H2 with residual O2 present in the jar to form water. An indicator, i.e. reduced Methylene blue is employed for verifying anaerobic condition in the jar, which remains colourless anaerobically, but turns blue on exposure to O2. The jar with the complete anaerobiosis is incubated at 37oC for 48 hours and growth of anaerobic organisms is noted.

21 Important Bacterial Pathogens and the Diseases ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

BACTERIAL PATHOGENS AND RELATED DISEASES Organism

Morphology

Important diseases

Gram Positive Cocci Staphylococcus aureus

Cocci in groups

Boils, abscesses, secondary infections bacteraemia, pneumonia, meningitis, conjunctivitis in newborns, food-poisoning

Staphylococcus saprophyticus

Cocci in groups

Urinary tract infections

Streptococcus pyogenes (Group A Streptococcus)

Cocci in chains

Sore throat, scarlet fever, bacteraemia, otitis media, meningitis, cellulitis, puerperal sepsis. Post-Streptococcal glomerulonephritis, and rheumatic fever leading to heart disease Contd...

Important Bacterial Pathogens and the Diseases 89 Contd... Viridans Streptococci

Cocci in chains

Bacterial endocarditis, bacteraemia, tooth decay, abscesses.

Streptococcus pneumoniae (Diplococcus pneumoniae)

Capsulated diplococci

Meningitis, lobar pneumonia, otitis media, pleurisity

Streptococcus faecalis (Group D Streptococci)

Cocci in chains and pairs

Urinary tract infections, wound and ulcer infections, septicaemia

Streptococcus agalactiae (Group B Streptococcus)

Cocci in chains and pairs

Septicaemia, Pneumonia, neonatal meningitis

Gram Positive Bacilli Bacillus anthracis

Large, spore forming, capsulated, bacilli which tend to form chains

Anthrax

Corynebacterium diphtheriae

Pleomorphic, non-motile rods, often seen joined at angles

Diphtheria of the throat and skin

Listeria monocytogenes

Small rods with tumbling motility at low temperatures

Meningitis, still-birth, bacteraemia

Clostridium botulinum

Non-motile Severe food-poisoning pleomorphic rods, with oval subterminal spores Contd...

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Contd... Clostridium tetani (Drum-stick bacillus)

Non-motile, long thin rods with oval subterminal spores

Tetanus

Clostridium perfringens (Clostridium welchii)

Non-motile, thick, brick-shaped rods, Spores are rarely Gas gangrene, septicaemia, seen food poisoning

Clostridium difficile

Non-motile, thick, Antibiotic-associated brick-shaped rods, diarrhoea and colitis Spores are rarely seen Gram Negative Rods

Haemophilus influenzae

Small, non-motile Acute respiratory infections, coccobacilli or meningitis, cellulitis, rods, pleomorphic, ear infections Usually capsulated

Haemophilus aegyptius

Small, non-motile rods or cocco bacilli

Haemophilus ducreyi

Small, non-motile Soft chancre rods often in pairs (Chancroid) or chains

Bordetella pertussis

Small, non-motile coccobacilli, may be capsulated

Whooping cough

Brucella species

Small, non-motile coccobacilli may show bipolar staining

Brucellosis (undulant fever)

Yersinia pestis

Non-motile, capsulated cocco-bacilli, showing bipolar staining

Bubonic, septicaemic and pneumonic plague

Infectious conjunctivitis (pink-eye), respiratory infections.

Contd...

Important Bacterial Pathogens and the Diseases 91 Contd... Yersinia pseudotuberculosis

Motile pleomorphic rods or coccobacilli showing bipolar staining

Acute mesenteric lymphadenitis, septicaemia

Yersinia enterocolitica

Motile pleomorphic rods or cocco bacilli showing bipolar staining

Gastroenteritis, peritonitis, bacteraemia, abscesses

Francisella tularensis

Small, non-motile, capsulated, pleomorphic rods or coccobacilli, showing bipolar staining

Skin infections, lymphadenitis, eye infections, lymphoid-like illness, and respiratory infections.

Bacteroides species

Non-motile small pleomorphic rods

Bedsores, abscesses, abdominal and pelvic infections, bacteraemia

Escherichia coli

Motile rods

Urinary infections, wound infections, bacteraemia, gastroenteritis

Klebsiella pneumoniae

Non-motile capsulated rods

Chest infections, urinary infections, wound infections bacteraemia, meningitis, endocarditis

Klebsiella rhinoscleromatis

Non-motile capsulated rods, with foam cells

Rhinoscleroma of the upper respiratory tract

Proteus species

Motile rods

Urinary infection, respiratory infections, ear and wound infectinos, burns infections (often hospital-acquired), septicaemia

Pseudomonas pseudomallei

Motile rods showing bipolar staining

Melioidosis (pneumoenteritis) Contd...

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Contd... Shigella species

Non-motile rods

Bacillary dysentery

Salmonella species

Motile rods

Enteric fever (typhoid and paratyphoid), food-poisoning, septicaemia, meningtis, bone infections, abscesses

Vibrio cholerae

Motile, slightly curved rods

Cholera

Vibrio parahaemolyticus

Motile, slightly curved rods

Gastroenteritis

Campylobacter jejuni/coli

Motile, curved spiral rods

Gastroenteritis

Legionella pneumophila

Non-motile rods Severe pneumonia Gram Negative Cocci

Neisseria meningitidis

Intracellular diplococci

Meningitis and septicaemia

Neisseria gonorrhoeae

Intracellular diplococci

Gonorrhoea, eye infection in newborns

Acid Fast Bacilli Mycobacterium tuberculosis

Non-motile bacilli

Tuberculosis

Mycobacterium ulcerans

Non-motile bacilli

Skin ulcers (Buruli ulcer)

Mycobacterium leprae

Non-moitle bacilli Leprosy often in groups (globi) Spirochaetes and Other Organisms

Treponema pallidum1

Motile delicate treponemes

Syphilis

Treponema pertenue1

Motile delicate treponemes

Yaws Contd...

Important Bacterial Pathogens and the Diseases 93 Contd... Treponema carateum1

Motile delicate treponemes

Pinta

Leptospira interrogans1

Motile thin leptospires with hooked ends

Epidemic spirochaetal jaundice (leptospirosis iceterohaemorrhagica), Weil’s disease, Canicola fever

Borrelia vincenti2

Motile borreliae Associated with Vincent’s (Gram negative) angina (Trench mouth), and tropical ulcer

Borrelia duttoni2

Motile borreliae

Tick-borne relapsing fever

Borrelia recurrentis2

Motile borreliae

Louse-borne relapsing fever

Spirillum minus2

Motile, small, Rat bite fever (Sodoku) rigid spirals with flagella

Bartonella bacilliformis2

Rod-shaped Oroya fever (Carrion’s organisms found disease), Verrugaperuana in red cells and reticuloendothelial cells

Rickettsia species3

Intracellular Typhus fevers (epidemic and organisms just endemic), trench fever, visible with the Q fever light microscope

Chlamydia species4

Inclusion bodies in epithelial cells

Calymmatobacterium granulomatis (Donovania granulomatis)2

Small pleomorphic Granulomatous disease rods found inside (Granuloma venereum) mononuclear cells

Trachoma, inclusion conjunctivitis, lymphogranuloma venereum, non-gonococcal urethritis

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Notes: 1. Treponemes and Leptospires are best seen using dark-field microscopy. 2. Borrelia, Bartonella, and Calymmatobacterium species are best seen in Giemsa stained preparations. Spirillum minus is Gram negative but can also be stained using Giemsa. 3. Rickettsiae can be stained using Giemsa stain but it is usual to diagnose rickettsial infections serologically. 4. Chlamydiae can be seen in Giemsa and iodine stained preparations.

22 Collection of Clinical Materials for Microbiological Investigations ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Microbiological laboratories aid the clinicians in many ways. First and fore most is the diagnosis of disease process and treatment of it. Hence care should be taken during collection of the specimens at bed sides. The following general rules must be observed while collecting a clinical sample. • The specimens should be from the suspected infective condition. • Sterile swabs and sterile containers should be used to avoid contamination. These materials can be supplied from the Microbiology department. • The sample should be in sufficient quantity to permit a thorough study. • It is always better to send the sample before administration of antibiotic, if not; the antibiotic used should be informed. • The specimens should be clearly labelled with patient’s name, date of collection, time of collection and hospital number. This

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A. SPECIMEN FOR BACTERIOLOGICAL INVESTIGATIONS 1. Blood for Culture The following apparatus are required, Sterile 10 ml syringe, 21 gauge needle, antiseptic solution to clean the skin, spirit lamp and Media for culture of different organisms. Organisms Media Quantity of Blood Salmonella 50 ml bile broth Cocci 50 ml glucose broth 10 ml Anaerobic organisms Thioglycollate broth 5 ml Brucella Tryptose soy broth Or Liver infusion broth 5 ml General precautions while collecting blood The skin should be cleaned properly with antiseptics. 2% tincture Iodine and later with 70% alcohol. The site is allowed to dry and blood should be collected in the following clinical conditions. 1. Enteric fever 2. Subacute bacterial endocarditis 3. Brucellosis 4. Any Septicaemias. • For enteric fever blood should be collected in the first week of theillness. • For septicaemias, 4 frequent samples should be collected at subsequent temperature in the same day. • The Samples of blood should be sent to the laboratory immediately. 2. Urine • Urine should be collected in sterile wide mouth containers.

Collection of Clinical Materials for Microbiological Investigations 97

Figure 22.1: Blood culture bottle with bile broth (For colour version, see Plate 6)

Figure 22.2: Centrifuge

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Figure 22.3: Sterile swab (For colour version, see Plate 6)

Figure 22.4: Throat swab collection

• In females: Midstream urine sample should be collected after cleaning the external genitalia with soap and water. In some occasions catherterisation should be done, but always avoid catherterisation since there is a chance of introducing microorganisms in to the bladder. • In males: The glans is cleanced with soap and water and midstream urine should be collected.

Collection of Clinical Materials for Microbiological Investigations 99 • Urine specimens should be brought to the laboratory immediately with-in one hour. If that is not possible they should be kept in the refrigerator for not more than 24 hours time. 3. Stool Samples Should be collected in sterile containers and immediately sent to the laboratory. 4. Rectal Swab This can be taken by introducing the sterile swab into the anus, and the swab should be replaced in a test tube. 5. Sputum The sputum should be collected in sterile screw capped bottles. 6. Throat Swab This is a sterile swab fitted to a wooden stick kept inside sterile test tube; outside fitted with a cotton plug. Make it a point always to send two swabs to the laboratory—One for smear and one for culture. The swab stick should be removed from the tube. Swab the back of the throat; without touching the other parts of the mouth. Replace the swab in the tube. Despatch it as soon as possible. 7. Nasal, Nasopharyngeal and Oral Swabs Nasal, nasopharyngeal and oral swabs should be collected in the same way as throat swabs. 8. Ear Swab Two swabs should be taken, one for the mycological examination another for the bacterial cultures. 9. CSF CSF should be collected(lumbar puncture) under sterile precautions 2-5 ml of it can be sent to the laboratory in sterile test tubes or screw capped bottle without any delay.

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10. Conjunctival Swab The swab should be taken before starting on any local antibiotic therapy; if started it should be washed with sterile saline and then the swab should be taken from the suspected area. 11. Cervical and Vaginal Swabs

Vaginal swab: The site should be cleaned with savalon and then the swab should be taken from the suspected site Cervical swab: Speculum should be used to visualize the cervix and cervical swab should be taken under sterile precautions and sent to the laboratory. 12. Urethral Swab and Prostatic Fluid Two samples should be sent to the laboratory, one for direct smear examination one for culture. Swabs should be sent to the laboratory immediately or they can be inoculated for Gonococci at the bed side. 13. Pus Pus from close abscess can be collected by sterile syringe and needle, by aspiration, afterwards sterile swab is used. 14. Serous Fluid Pus from close abscess can be collected by sterile syringe and needle, by aspiration, afterwards sterile swab is used. Biopsy and autopsy specimens should be sent to the laboratory in sterile containers. B. SPECIMENS FOR ACID FAST BACILLI 1. Sputum: Over night sputum in a wide mouthed sterile container should be sent to the laboratory. 2. Urine for tubercle bacilli: In a clean bottle 24 hours collected sample should be sent or else entire early morning sample should be sent.

Collection of Clinical Materials for Microbiological Investigations 101 3. Laryngeal swab and gastric lavage: When very little sputum is present in children who swallow sputum, materials should be obtained either by laryngeal swab or by gastric lavage. Laryngeal swab: Two consecutive swabs should be taken from each patient. Gastric lavage: Fasting stomach contents may be aspirated with Ryle’s tube in the morning. The sample should be sent within an hour of collection. 4. Pus from cold abscess: Small quantity of pus can be sent in a sterile pencillin bottle. 5. Pleural and peritoneal fluids: About 100 cc. of fluid should be sent in a sterile flash to which citrate is added prior to the collection of fluids. 6. CSF: About 5 cc of CSF can be sent in a sterile screw capped bottle. 7. Faeces: About 5 grams of faeces should be sent in a clean bottle. 8. Tissues: Biopsy or autopsy specimens should be sent in sterile dry containers. 9. Tissues smear for lepra bacilli: Two smears should be sent for examination nasal swabs can also be sent. C. SPECIMENS FOR ANAEROBIC CULTURE 1. Pus: Swabs should be taken from the deeper parts of the wounds and abscesses with sterile stick swab. The stick should be broken at the tip and swab should be kept in thioglycollate broth or RCM and sent to the Laboratory immediately. 2. Blood: About 5 ml of blood should be sent in thioglycollate broth. 3. Urine: Midstream urine should be sent in thioglycollate broth. 4. Prostatic fluid: Prostatic fluid should be collected under sterile condition into the thioglycollate broth. 5. Serous fluid: Pleural fluid under aseptic condition should be collected into the thioglycollate broth and sent to the laboratory. D. BLOOD FOR SEROLOGICAL TESTS Dry sterile syringe and sterile needle should be used. The blood should be sent in sterile screw capped bottles or sterile test tubes. The sample must be accompained with requisition form with all

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relevant informations. Preferably two samples of blood should be sent one immediately after the admission and another a week later. This is necessary to study the rise in antibody titre. In Widal, VDRL, Brucella agglutination test, RA latex fixation test, Paul Bunnell test, CRP test, and ASO test, send 5 ml of clotted blood in sterile test tubes or screw capped bottles. E. MATERIALS FOR MYCOLOGICAL INVESTIGATIONS 1. Skin Scrapings: The affected area is swabbed with sterile gauze soaked in 70% alcohol and allowed to dry. The lesion is scrapped with a sterile surgical blade at the edge of the lesion. These scrapings should be kept inside a sterile petridish or a sterile paper packet. This should always be accompained with a requisition form furnishing available data. 2. Hairs: The unhealthy hairs present at the site of lesion should be plucked with sterile tweezers and kept in a sterile petridish and sent to the laboratory. 3. Nail: A piece of the affected nail and the scrapings from the under surface of nails are sent to the laboratory. 4. Materials from systemic fungal infections: Blood, spinal fluid, sputum, exudate from the draining sinuses and abscesses, sternal marrow, scrapings from the edges of ulcers and abscesses and biopsy tissues are placed in sterile vials or petridishes or test tubes and sent to the laboratory. 5. Swabs from mouth and vagina: The swabs taken from the mucosal surface for monilliasis are placed in a sterile test tube and sent to the laboratory. Biopsy specimens: These are sent in sterile container for culture and animal inoculation. F. SPECIMENS FOR PARASITOLOGICAL EXAMINATION 1. Faeces: For Examination of parasites, ova and freshly voided faeces must be sent in a clean bottle. 2. Blood for microfilaria: At least 2 cc of midnight sample of blood should be sent with the anitcoagulant added to it.

Collection of Clinical Materials for Microbiological Investigations 103 3. Other specimens: Materials like splenic puncture, liver puncture, pus from liver abscess, CSF, urine rectal swabs and blood may also be sent for parasitological examination. Note: Stool for culture must be sent in a sterile bottle. G. SPECIMENS FOR VIROLOGICAL INVESTIGATION The diagnostic virology is entirely dependent on the correct selection of clinical specimens and their collection and transport to the laboratory under appropriate conditions. Specimen should be collected early during the course of the illness as successful isolation may be less in the later stages. All specimens should be colleted in sterile containers. Each specimen should be accompanied with requisition, from containing patient details like name, age sex, ward and hospital number with brief history of nature of specimen, and date of collection and examination required. Most viruses are unstable above 40 oC, hence specimens should be sent immediately, preferably kept in chilled packet in ice to the laboratory. For isolation of certain viruses the specimens should be collected in a transport medium. The common transport medium used for virological specimens is Hank’s balanced salt solution + 10% bovine albumin + Sodium carbonate + 50 units of penicillin + 50 mg of Sterptomycin. The Specimens that can be transported in this medium are swabs, biopsy and autopsy materials. The seriological tests can be conducted where the isolation of viruses is impractical, and also to find out significance of the isolation of certain viruses. Specimens for serological tests should not be frozen, but should reach the laboratory within 24 hours of collection. For serological procedure two samples of clotted blood should be sent once during the acute illness and another 14 days after the 1st sample. They should be sent in sterile screw capped bottles. a. Diseases of central nervous system 1. Faeces: The specimen should be collected from the acute stage upto IInd week 5-10 grams of Faeces should be collected in a

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sterile screw capped bottle. The contents should fill only 3rd of the bottle. Freezing is not necessary as certain viruses are stable at normal room temperature. CSF: 5 ml collected during the acute stage of the illness. The specimen should be chilled by keeping the container in ice cubes and sent to the laboratory. Throat swabs: The specimen should be collected during the first few days of illness. After swabing it is immediately broken into a sterile screw capped bottle containing 5 ml of transport medium. Neuropsy specimens: Several grams of the brain substance can be sent to the laboratory in the frozen state. Blood: 5 ml of heparinised blood should be collected during first few days of suspected infection. Saliva: It is collected for suspected rabies cases. Paired sera are collected for serological tests.

b. Disease of the Skin Macules or papules should be scrapped with scalpel. Smears should be made on several glass slides. Vesicular fluid should be collected into capillary tubes. These should be kept in screw capped bottles. After removing the vesicular fluid scrapings from the base of the vesicles should be taken and smeared on slides for microscopic examination. The specimens may be dangerous and proper care should be taken while handling them. c. Disease of the Eye Sterile swabs with wooden stick should be used. After collecting swabs from the diseased part of the eye, the tip of the swabs should be cut and the swab should be kept in a screw capped bottle containing 5 ml of transport medium; at the same time several smears can be prepared on glass slides. d. Diseases of the Respiratory Tract The specimens collected in the respiratory viral infections are mouth garglings, Throat swabs and nasopharynegeal swabs. They should

Collection of Clinical Materials for Microbiological Investigations 105 be collected in the first 3 days after the illness. After taking the specimens they should be placed in screw capped bottles containing 5 ml of transport medium and sent to the laboratory. H. MATERIALS FOR STERILITY TESTS Suture materials, gauze pieces, cotton, catgut, can be sent in sterile bottle or test tubes.

Spore paper is supplied by the Microbiology laboratory. This should be kept in the centre of the articles to be sterilized before autoclaving. The same to be sent to the laboratory to know whether the sterilization is proper or not. Once in every 15 days, swabs may be collected from theatre, equipment and sent to the laboratory for testing anaerobic spore bearing organisms.

23 Biochemical Tests and Identification of Bacteria ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

After the successful cultivation of bacteria in the laboratory, the next step is identification of the genus and species of the cultivated organism. For the identification individual bacteria various methods are available. They include staining techniques, biochemical tests, serological tests, animal pathogenecity tests and nucleic acid techniques. Among these, biochemical tests are very useful for the routine identification of pathogenic bacteria. The Validity of the identification of an unknown bacterial culture by its reaction in a range of biochemical tests depends on the use of a pure culture of the bacteria for inoculation of the test media. Single, well separated colonies grown on a non-selective culture medium should be used as inoculum for the tests. Common Biochemical Test Used to Differentiate Bacteria 1. 2. 3. 4. 5.

Catalase test Oxidase test Sugar fermentation test O/F test Citrate utilisation test

Biochemical Tests and Identification of Bacteria 107 6. 7. 8. 9. 10. 11. 12. 13. 14.

Bile solubility tests Indole production test Coagulase test Voges-Proskauer test Methyl red test Urease test Nitrate reduction test Coagulase test H2S production test.

24 O/F Test (Hugh and Leifson's Test) ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

This test is used to distinguish between aerobic and anaerobic break down of Carbohydrate by Bacteria. A semisolid medium containing glucose and pH indicator is used for the test. if acid is produced only at the surface of the medium, where conditions are aerobic, the attack on the sugar is oxidative if acid is found throughout the tube, including the lower layers where conditions are anaerobic the breakdown is fermentative (Fig. 24.1). Requirements 1. Medium Peptone - 2 gm NaCl - 5 gm K2HPO4 - 0.3 gm Bromothymol blue (1% aqueous solution) : 3 ml Agar - 3 gm Water - 1 litre The pH is adjusted to 7.1 before adding bromothymol blue. Autoclave the medium.

O/F Test (Hugh and Leifson's Test) 109

Figure 24.1: O/F test

The medium is then tubed to a depth of about 4 cm. 2. Straight wire 3. Sterile melted petroleum jelly 4. liquid culture (test organism). Method • Two tubes of medium are taken. • Inoculate the medium by stabbing. • One tube is promptly sealed with a layer of sterile melted petroleum jelly to a depth of 5-10 mm. • One tube without sealing. • Incubate both tubes. Results

Open tube

Sealed tube

Oxidation (e.g. : Acinetobacter) Fermentation (e.g. : Esch.coli) No action on Carbohydrate (e.g. : Alcaligenes faecalis)

Yellow Yellow

Green Yellow

Green

Green

Note: This medium can also be used for detecting gas production and motility.

25 Catalase Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE This test demonstrates the presence of catalase, an enzyme that catalyses the release of oxygen from hydrogen peroxide. Hydrogen peroxide is a by-product of aerobic respiration and is a toxic substance for bacterial cell. Aerobic bacteria produce catalase to breakdown hydrogen perioxide into water and oxygen. This can be demonstrated in vitro by catalase test. Requirements 1. Hydrogen peroxide solution (10 vol). 2. Platinum wire loop or Plastic loop or sealed capillary tube. 3. Pure growth bacteria. Method • Take H2O2 solution in a clean test tube. • Pick up culture to be tested from a nutrient agar slope with a clean sterile platinum loop or sealed cappillary tube. • Insert the wire loop into H2O2 Solution.

Catalase Test 111 Result • Immediate Production of gas bubbles from the surface of the solid culture material – Positive e.g. Staphylococcus spp. • No bubbles – Negative e.g. Streptococci spp. Note: A false positive reaction may be obtained if the culture medium contains catalase (e.g. Blood agar) or if an iron wire loop is used.

26 Oxidase Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE This test depends on the presence in bacteria of certain oxidases that will catalyse the transport of electrons between electron donors in the bacteria and a redox dye – tetra methyl-p- phenylene-diamine. The dye is reduced to a deep purple colour. This test is very useful in screening of Neisseria, Aeromonas, Vibrio, Campylobacter and Pseudomonas, which give positive reactions and for excluding the enterobacteriaceae which give negative reactions. Requirements 1. 1% solution of tetramethyl – P-phenylene diamine dihydrochloride 2. Whatman’s no.1 filter paper. 3. Glass rod/platinum wire loop. Methods The test can be performed by three methods: 1. Plate method

Oxidase Test 113 2. Dry filter paper method 3. Wet filter paper method. 1. Plate Method • Cultures are made on suitable solid culture medium. (e.g.: nutrient agar) • A freshly prepared 1% solution of tetramethyl–p-phenylenediamine dihydrochloride is poured on to the plate. • Decant the solution. Results • Colonies rapidly develop a purple colour–Oxidase positive • Colonies do not change their colour–Oxidase negative 2. Dry filter paper method • Strips of Whatman’s no.1 filter paper are soaked in a freshly prepared solution of tetramethyl–p-phenylene-diamine dihydrochloride. • After draining for about 30 seconds the strips are freeze dried and stored in a dark bottle tightly sealed with a screw cap. The papers will have a light purple tint. • A strip is laid in a petridish and moistend with sterile distilled water. • The colony to be tested is picked up with a platinum wire loop and smeared over the paper. Results Intense deep-purple colour on the paper develops within 5-10 seconds positive oxidase. Absence of colourisation or colouration after 60 seconds negative oxidase (Fig. 26.1). 3. Wet Filter Paper Method • Strips of Whatman’s no.1 filter paper are soaked in a freshly prepared solution of oxidase reagent.

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Figure 26.1: Oxidase test (For colour version, see Plate 7)

• Place the paper in a petridish. • Smear the paper with the culture to be tested with a platinum wire loop. • Result same as for the dry filter paper method. Example Oxidase Positive Bacteria • Pseudomonas • Neisseria • Vibrio.

Example: Oxidase Negative • Esch. coli • Salmonella • Shigella.

27 Sugar Fermentation Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE Many bacteria are able to ferment a variety of carbohydrates or related compouds. They are broken down to acids like lactic acid, pyruvic acid, or in to gases like CO2 or hydrogen. The fermentation range of a particular bacterial species is usually constant. By performing a battery of sugar fermentation tests we can identify a species. In a routine laboratory various carbohydrates like glucose, lactose, maltose, inulin, salicin, esculin, mannitol, sucrose, etc. are used. If the organism ferments a given carbohydrate the pH of the medium turns into acidic and this can be detected by using a suitable pH indicator. Gas production can be detected by using an inverted Durham’s tube in the medium (Fig. 27.1). Requirments 1. Medium 2. Sugar

- Peptone water/Serum peptone water/Serum agar. - Glucose/Fructose/Mannose/Galactose/ Sucrose/Maltose/ Lactose/Salicin, etc. 3. Suitable indicators - Andrade’s indicator - Bromocresol purple

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Figure 27.1: Sugar fermentation test (phenol red indicator) showing negative, acid formation, and acid and gas formation (For colour version, see Plate 7)

- Phenol red - Bromothymol blue 4. Inverted Durham’s tube Method 1. Peptone water fermentation test (common method) 2. Serum peptone water fermentation test. This media are suitable for more exacting organism like Corynebacterium diphtheriae. 3. Serum agar fermentation test: This media are recommended for organisms such as Meningococci and Gonococci that grow poorly in liquid media. The commonest nutrient medium for fermentation tests is peptone water. The nutrient medium is prepared and indicater added. • Each carbohydrate is incorporated in to peptone water to a concentration of 1%. • Add pH indicator (Commonly used indicator is Andrade’s indicator. It is made by adding NaOH 1 mol/litre to a 0.5% solution of acid fuchsin until the colour just becomes yellow. It is used at a a final concentration of 0.005% in the medium. It turns dark reddish pink at about pH 5.5 (Fig. 27.2).

Sugar Fermentation Test 117

Figure 27.2: Sugar fermentation test (Andrade’s indicator) showing negative reaction and acid formation

• The Medium is sterlized after distributing into sugar tubes with a small inverted glass tube called Durham’s tube. • Before inoculation, confirm the absence of bubbles gas from Durham’s tube in liquid media. • Inoculate the sugar media with a drop or loopful of liquid culture. • Incubate at 37 oC for 24 hours. Result Pink colour with gas bubble in the Durham’s tube

Carbohydrate fermented with acid and gas (+) (e.g. Esch.coli, Klebsiella)

Pink colour with no gas bubbles in the Durham’s tube

Carbohydrate is fermented without gas production (+) (e.g. Salmonella Shigella, Vibro)

No color change or yellow colour Carbohydrate not fermented (-) and no gas in Durham’s tube e.g. Pseudomonas, Alkaligenes

28 Nitrate Reduction Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE This test detects the presence of enzyme nitrate reductase which causes the reduction of nitrate to nitrite in the presence of a suitable electron donor. Almost all Enterobacteriaceae members reduce nitrate. Nitrate reduction can be detected by an appropriate colorimetric reagent. Requirement 1. Medium Potassium nitrate - 0.2 gm Peptone - 5 gm Distilled water - 1litre Tube in 5 ml amounts and autoclave at 121oC for 15 min. 2. Test reagent Solution A - Dissolve 8 gm sulphanilic acid in 1 litre of acetic acid 5 mol/lit Solution B - Dissolve 5.0 gm of α-naphthylamine in litre of acetic acid 5 mol /litre immediately before use, mix equal volume of solution A and B to give the test reagent.

Nitrate Reduction Test 119 Method • Inoculate the medium with test organism • Incubate at 37 oC for 96 hours. • Add 0.1 ml of the test reagent to the test culture. Result A red color within a few minutes - Nitrate reduction positive e.g.Esch.coli No.red color - Nitrate reduction negative e.g. Acinetobacter.

29 Hydrogen Sulphide Production Test

○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE Some gram negative bacillii produce H2S by breaking down sulphur containing amino acids like cysteine or cystine. Hydrogen sulphide is demonstrated by its ability to form black insoluble sulphides. Method 1 There are different methods and media are used for detection H2S production. List of media used for H2S detection 1. 2. 3. 4. 5. 6.

Bismuth sulfite agar. Triple sugar iron agar (TSI). Lysine iron agar. Lead acetate agar. Kigler iron agar. Deoxycholate citrate agar.

Hydrogen Sulphide Production Test 121 Method 2 Filter paper method. • Prepare a Whatman No.1 filter paper impregnated with lead acetate (10% solution) • Expose the filter paper above culture by placing filter paper between the cotton plug and test tube. Result Blackening of the lead paper

H2 S positive (Citrobacter, Salmonella

No blackening of the lead acetate paper

H2S negative (e.g. Esch. coli, Klebsiella)

30 Urease Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE Some bacteria, particularly those growing naturally in an environment exposed to urine, may decompose urea by enzyme urease and liberate ammonia Urea →Ammonia + CO2 + Water. The production of the enzyme urease is detected by growing the organisms in the presence of urea and testing for alkali (NH3) production by means of a suitbale pH indicator (Fig. 30.1). Requirements 1. Medium – Christensen’s urea agar. Peptone - 1 gm NaCl - 5 gm K2HPO4 - 2 gm Phenol Red (1 in – 500 aqueous solution) - 6 ml Agar - 20 gm Distilled water - 1 litre

Urease Test 123

Figure 30.1: Urease test (For colour version, see Plate 8)

Glucose 10% solution - 10 ml Urea 20% solution - 100 ml Sterilise the glucose and urea solutions by filteration. Prepare the basal medium without glucose or urea, adjust the pH to 6.8-6.9, sterilised by autoclaving, cool to about 50oC, add the glucose and urea and tube the medium as deep slopes. Method • Inoculate the medium heavily over the entire slope surface with the test organism culture. • Incubate at 37 oC Overnight incubation. Result Colour of the medium changes to purple pink

Urease positive (e.g. Proteus, Klebsiella)

No colour change

Urease negative (e.g. Esch.coli)

31 Citrate Utilisation Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE Some bacteria can utilise simple organic salts like sodium citrate or sodium acetate as the sole source of carbon and energy source for growth and an ammonium salt as the sole source of nitrogen. During such utilisation CO2 is liberated which makes the medium alkaline. Koser’s liquid citrate medium or Simmon’s citrate agar may be used. Requirements 1. Medium: Simmon’s citrate agar medium. (This is a modification of Koser’s medium with agar and an indicator added) (Fig. 31.1). Sodium chloride 5 gm Magnesium sulphate 0.2 gm NH4H2PO4 1 gm 1 gm KH2PO4 Sodium citrate 5 gm Distilled water 1 litre Agar 20 gm Bromothymol blue 0.2% 40 ml.

Citrate Utilisation Test 125

Figure 31.1: Simmons’s citrate media (For colour version, see Plate 8)

Adjust the pH to 6.8. Sterilise by autoclaving at 121oC for 15 min. Allow to set as slopes in test tubes. Method • Inoculate the slope with a wire loop from a saline suspension of the organism to be tested. • Incubate at 37oC for 96 hours. Result Citrate utilised

Growth with deep blue slant (e.g. Klebsiella, Proteus). Citrate not utilised Original green colour and No growth (e.g. Esch.coli, Shigella spp.

32 Voges–Proskauer Test (VP Test)

○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE Some bacteria ferment carbohydrates with the production of acetyl methyl carbinol or its reduction product 2, 3 butylene glycol. This breakdown product when reacts with Alpha-Naphthol in alcohol in the presence of alkali gives a cherry red colour. This test is usually done in conjunction with the methyl red test since the production of acetyl methyl carbinol or butylene glycol usually results in insufficient acid accumulation during fermentation to give a methyl red positive reaction. An organism of the Enterobacteriaceae family is usually either MR positive and VP negative or MR negative and VP Positive (Fig. 32.1). Requirements 1. Medium : Glucose phosphate peptone water (as for the MR Test) 2. 40% potassium hydroxide 3. 5% solution of alpha –naphthol in absolute ethanol

Voges-Proskauer Test (VP Test) 127

Figure 32.1: VP test (For colour version, see Plate 9)

Method • • • •

Inoculate the liquid medium Incubate at 37% for 48 hours. Add 1ml of 40% potassium hydroxide and 3 ml of alpha naphthol. Shake well and keep it in the rack for 10 minutes

Result Pink or cherry red colour Yellow colour

VP Positive (e.g. Klebsiella) VP negative (e.g. Escherichia coli)

33 Methyl Red Test (MR Test)

○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE This test is used to detect the production of large amount of acid during the fermentation of glucose and the maintenance of conditions such that the pH of an old culture is sustained below a value of about 4.5. The acidity can be determined by adding a few drops of methyl red solution, as indicator (Fig. 33.1). Requirements 1. Medium : Glucose phosphate peptone water Peptone : 5 gm K2HPO4 : 5 gm Water : 1 litre Glucose, 10% solution : 50 ml (Sterilised separately) Dissolve the peptone and phosphate, adjust the pH to 7.6, Filter, dispense in 5 ml amounts and sterilise at 121oC for 15 min. Sterilise glucose solution by filteration and add 0.25 ml to each tube final concentration (0.5%). 2. Methyl red indicator solution:

Methyl Red Test (MR Test)

129

Figure 33.1: MR test (For colour version, see Plate 9)

Methyl red : 0.1 gm Ethanol : 300 ml Distilled water : 200 ml Method • Inoculate the liquid medium lightly from a young agar slope culture. • Incubate at 37oC for 48 hours. • Add five drops of the methyl red reagent. • Mix and read immediately. • Positive tests are bright red. • Negative tests are yellow. Result Bright red Yellow

Positive (e.g. Escherichia coli) Negative (e.g. Klebsiella spp)

34 Indole Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE This test demonstrates the ability of certain bacteria to decompose the amino acid Tryptophan to Indole. When a bacterium is grown in medium containing Tryptophan (Such as peptone water), it breaks down the Tryptophan producing indole. This indole when combined with a compound called para dimethyl amino benzaldehyde (added in the form of Kovacs reagent) gives a pink coloured compound called Rose Indole (Figs 34.1 and 34.2). Requirements 1. Medium : Peptone water 2. Kovac’s reagent : Amyl or iso amyl alcohol : 150 ml P-Dimethyl-amino benzaldehyde : 10 gm Hydrochloric acid (Concentrated) : 1 litre Dissolve the aldehyde in the alcohol and slowly add the acid. Prepare in small quantities and store in the refrigerator. Shake gently before use.

Indole Test 131

Figure 34.1: Indole test (tube method)

Figure 34.2: Indole spot test

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Method • Inoculate peptone water medium with test organism and incubate for 48 hours at 37oC. • Add 0.5 ml Kovac’s reagent and shake gently. • A red color in the alcohol layer indicates a positive reaction. Result Pink colour No colour

Indole positive bacteria – Escherichia coli Indole negative bacteria – Klebsiella spp.

Note The test can also be conducted by inserting a paper (dipped in the reagent) at the mouth of the test tube containing over night peptone water culture of the test organism. A pink colour develops in tip of the filter paper on indole production.

35 Bile Solubility Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

This test is useful to differentiate Streptococcus pneumoniae (Pneumococcus) from viridans Streptococci. Pneumococcus Colonies are soluble in bile and bile salts and viridans Streptococci are insoluble. PRINCIPLE A heavy inoculum of the test organism is emulsified in physiological saline to give a turbid suspension. The bile salt sodium deoxycholate is then added. The bile salt dissolves Pneumococci as shown by a clearing of the turbidity within 10-15 minutes. Viridans Streptococci are not dissolved and therefore there is no clearing of the turbidity. Requirements 1. 2. 3. 4. 5.

Sodium deoxycholate (100 gm/lit, 10% W/V) Physiological saline (sodium chloride, 8.5 gm/lit) Test tubes Wire loop Distilled water.

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Procedure • Emulsify several colonies of the test organism in a tube containing 2 ml of sterile physiological saline, to give a turbid suspension. • Divide the organism suspension between two tubes. • To one tube, add 2 drops of the sodium deoxycholate reagent and mix. • To the other tube, add 2 drops of sterile distilled water and mix. • Leave both tubes for 10-15 minutes. • Look for a clearing of turbidity in the tube containing the sodium deoxycholate. Result Clearing of turbidity Test organism may be Pneumococci No clearing of turbidity Test organism is probably not Pneumococci Note There should not be clearing of turbidity in the tube to which distilled water is added. If there is tubidity clearing, repeat the test. Test can be performed along with positive and negative controls. Positive control – Streptococcus pneumoniae Negative control – Streptococcus faecalis

36 Coagulase Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

This test is used to differentiate Staphylococcus aureus which produces the enzyme coagulase, from Staph. epidermidis and Staph. saprophyticus which do not produce coagulase. PRINCIPLE Coagulase is an enzyme produced by Staphylococcus aureus. It converts fibrinogen to fibrin and causes clotting of plasma. Two types of coagulase are produced by most strains of Staph aureus. 1. Free coagulase 2. Bound coagulase Free coagulase converts fibrinogen to fibrin by activity a coagulase-Reacting factor present in plasma. Free coagulase is detected by the appearance of a fibrin clot in the tube coagulase test (Fig. 36.1). Bound coagulase (Clumping factor) converts fibrinogen to fibrin without the help of coagulase reacting factor. It can be detected by the clumping of bacterial cells in the slide coagulase test.

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Figure 36.1: Tube coagulase test

Procedure and Results of Slide Test (to detect bound coagulase) • Place a drop of saline on a slide. • Place a drop of saline on second slide also. • Emulsify a colony of the test organism in each of the drops to make two thick suspensions. • Add a drop of undiluted plasma to one of the suspensions, and mix gently. Look for clumping of the organisms within 10 seconds. • No plasma is added to the second suspension. This is used to differentiate any granular appearance of the organism from true coagulase clumping. Result Clumping within 10 seconds

Coagulase positive Staphylococci

No clumping within 10 seconds

Bound coagulase absent.

Tube Test: (to detect free coagulase) • Dilute the plasma 1 in 10 in physiological saline (mix 0.2 ml of plasma with 1.8 ml of Saline).

Coagulase Test 137 • • • • •

Take 3 small test tubes and label. T = Test organism P = Positive control N = Negative control Pick up a colony of test organism and inoculate in to the ‘T’ test tube. • Inoculate Staph.aureus colony in to ‘P’ test tube. • Incubate the 3 tubes at 37 oC. Examine for Clotting after 1 hour. If no clotting has occured, examine at 30 minute intervals for upto six hour. • When, looking for clotting, gently tilt each tube. If there is clotting a jelly like mass is formed. Result Fibrin clot No fibrin clot

Staphylococcus aureus No free coagulase produced.

37 Antimicrobial Susceptibility Testing ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Antibiotic susceptibility testing has become a very essential step for the proper treatment of infectious diseases. It is used. 1 To guide the clinician in selecting the best antimicrobial agent. 2. To accumulate epidemiological information on the resistance of microorganisms of public health importance. The choice of drugs used in a routine antibiogram is governed by various considerations since only a few antimicrobial agents can be tested (Fig. 37.1). Table 37.1 suggests the drugs to be tested in various situations. The drugs in Table 37.1 are divided into two sets. Set 1 includes drugs that are available in most hospitals and for which routine testing should be carried out for every strain. Tests for drugs in set 2 are to be performed only at the special request of the physician, or when the causative organism is resistant to the firstchoice drugs, or when other reasons (allergy to a drug, or its unavailability) make further testing justified.

Antimicrobial Susceptibility Testing 139 Table 37.1: Basic sets of drugs for routine susceptibility tests

Set 1

Set 2

Staphylococcus

Benzyl penicillin Oxacillin Erythromycin Tetracycline Chloramphenicol

Gentamicin Amikacin Co-trimoxazole Clindamycin

Enterobacteriaceae Urinary

Sulfonamide Trimethoprim Co-trimoxazole Ampicillin Nitrofurantoin Nalidixic acid Tetracycline

Norfloxacin Chloramphenicol Gentamicin

Blood and tissues

Ampicillin Chloramphenicol Cotrimoxazole Tetracycline Gentamicin

Cefuroxime Ceftriaxone Ciprofloxacin Piperacillin Amikacin

Pseudomonas aeruginosa

Piperacillin Gentamicin Tobramycin

Amikacin

Figure 37.1: Antibiogram

Antimicrobial susceptibility tests measure the ability of an antibiotic or other antimicrobial agent to inhibit bacterial growth in vitro. This ability may be estimated by either the dilution method or the diffusion method. The common method used in diagnostic microbiology laboratories is modified Kirby-Bauer method.

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MODIFIED KIRBY-BAUER METHOD Requirements

Mueller-Hinton Agar 1. Mueller-Hinton agar should be prepared from a dehydrated base according to the manufacturer’s recommendations. The medium should be such that with standard strains, zone sizes within the acceptable limits are produced. It is important not to overheat the medium. 2. Cool the medium to 45-50oC and pour into plates. Allow to set on a level surface, to a depth of approximately 4 mm. A-9 cm diameter plate requires approximately 25 ml of the medium. 3. When the agar has solidified, dry the plates for immediate use for 10-30 minutes at 36oC by placing them in an upright position in the incubator with the lids tilted. 4. Any unused plates may be stored in a plastic bag, which should be sealed and placed in a refrigerator. Plates stored in this way can be kept for two weeks. 5. In order to ensure that the zone diameters are sufficiently reliable for testing susceptibility to sulfonamides and co-trimoxazole, the Mueller-Hinton agar must have low concentrations of the inhibitors thymidine and thymine. Each new lot of MuellerHinton agar should therefore be tested with a control strain of Enterococcus faecalis (ATCC 29212 or 33186) and a disc of cotrimoxazole. A satisfactory lot of medium will give a distinct inhibition zone of 20 mm or more that is essentially free of hazy growth or fine colonies. 6. For testing the susceptibility of fastidious organisms, 5% blood should be added to the Mueller-Hinton agar base.

Antibiotic Discs Any commercially available discs with the proper diameter and potency can be used. Stocks of antibiotic discs should preferably be kept at 20oC, or the freezer compartment of a home refrigerator is convenient. A small working supply of discs can be kept in the refrigerator for up to one month. On removal from the refrigerator,

Antimicrobial Susceptibility Testing 141 the containers should be left at room temperature for about one hour to allow the temperature to equilibrate. This procedure reduces the amount of condensation that occurs when warm air reaches the cold container.

Turbidity Standard Prepare the turbidity standard by pouring 0.6 ml of a 1% (10 gm/L) solution of barium chloride dihydrate into a 100 ml graduated cylinder, and filling to 100 ml with 1% (10 ml/L) sulphuric acid. The turbidity standard solution should be placed in a tube identical to the one used for the broth sample. It can be stored in the dark at room temperature for six months, provided it is sealed to prevent evaporation.

Swabs A supply of cotton wool swabs on wooden applicator sticks should be prepared. These can be sterilized in tins, culture tubes, or on paper, either in the autoclave or by hot air oven. Method To prepare the inoculum from the primary culture plate, touch with a loop the tops of each of 3-5 colonies, of similar appearance, of the organism to be tested. When the inoculum has to be made from a pure culture, a loopful of confluent growth is similarly suspended in saline. Inoculum from colonies of streptococci cannot be made by emulsification. Hence, with streptococci, after inoculation the culture tubes are incubated for 4-6 hours to get uniform turbidity which should be matched with the turbidity standards. Compare the tube with the turbidity standard and adjust the density of the test suspension to that of the standard by adding more bacteria or more sterile saline. Proper adjustment to the turbidity of the inoculum is essential to ensure that the resulting lawn of growth is confluent or almost confluent.

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Inoculate the plates by dipping a sterile swab into the inoculum. Remove excess inoculum by pressing and rotating the swabs firmly against the side of the tube above the level of the liquid. Streak the swab all over the surface of the medium three times, rotating the plate through an angle of 60o after each application. Finally, pass the swab round the edge of the agar surface. Leave the inoculum to dry for a few minutes at room temperature with the lid closed. The antibiotic discs may be placed on the inoculated plates using a pair of sterile forceps. A sterile needle tip may also be used to place the antibiotic discs on the plate. Alternatively, an antibiotic disc dispenser can be used to apply the discs to the inoculated plate. A maximum of seven discs can be placed on a 9-10 cm diameter plate. Six discs may be spaced evenly, approximately 15 mm from the edge of the plate, and one disc placed in the centre of the plate. Each disc should be pressed down gently to ensure even contact with the medium. The plates should be placed in an incubator at 35oC within 30 minutes of preparation. Temperatures above 35oC invalidate the results for oxacillin/ methicillin. Do not incubate in an atmosphere of carbon dioxide. After overnight incubation, the diameter of each zone (including the diameter of the disc) should be measured and recorded in mm. The results should then be interpreted according to the critical diameters by comparing with standard tables (Table 37.2). The measurements can be made with a ruler on the under surface of the plate without opening the lid. The end-point of inhibition is judged by the naked eye at the edge where the growth starts, but there are three exceptions: 1. With sulfonamides and co-trimoxazole, slight growth occurs within the inhibition zone; such growth should be ignored. 2. When β-lactamase producing staphylococci are tested against penicillin, zones of inhibition are produced with a heaped-up, clearly defined edge; these are readily recognizable when compared with the sensitive control and regardless of the size of the zone of inhibition, they should be reported as resistant.

Antimicrobial Susceptibility Testing 143 3. Certain Proteus spp. may swarm into the area of inhibition around some antibiotics, but the zone of inhibition is usually clearly outlined and the thin layer of swarming growth should be ignored. Results The result of the susceptibility test, as reported to the clinician, is the classification of the microorganism in one of two or more categories of susceptibility. The simplest system comprises only two categories, susceptible and resistant. This classification, although offering many advantages for statistical and epidemiological purposes, is too inflexible for the clinician to use. Therefore, a three-category classification is often adopted. The Kirby-Bauer method and its modifications recognize three categories of susceptibility and it is important that both the clinicians and the laboratory workers understand the exact definitions and the clinical significance of these categories. Susceptible: An organism is called "susceptible" to a drug when the infection caused by it is likely to respond to treatment with this drug, at the recommended dosage. Intermediate susceptibility covers two situations. It is applicable to strains that are "moderately susceptible" to an antibiotic that can be used for treatment at a higher dosage because of its low toxicity or because the antibiotic is concentrated at the focus of infection (e.g. urine). The term also applies to those strains that are susceptible to a more toxic antibiotic that cannot be used at a higher dosage. In this situation the intermediate category serves as a buffer zone between susceptible and resistant.

Resistant: This term implies that the organism is expected not to respond to a given drug, irrespective of the dosage and the location of the infection. For testing the response of staphylococci to benzylpenicillin, only the categories ‘susceptible’ and ‘resistant’ (corresponding to the production of b-lactamase) are recognized. Staphylococci that are resistant to methicillin or oxacillin are also resistant to other penicillins or cephalosporins even though they show a zone of inhibition against these drugs.

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Zone diameters, to the nearest whole mm for various antimicrobial agents with disc content specified for each one for interpretation as susceptible, intermediate and resistant are given in Table 37.2. Table 37.2: Quality control – Susceptibility of control strains

Zone diameter of inhibition (mm) Antibiotic

Disc potency S.aureus (IU or µg) (ATCC 25923)

Amikacin 30 Ampicillin 10 Ceftriaxone 30 Cephalothin 30 Chloramphenicol 30 Ciprofloxacin 5 Clindamycin 2 Erythromicin 15 Gentamicin 10 Nalidixic acid 30 Nitrofurantoin 300 Norfloxacin 10 Oxacillin 1 Penicillin G 10 Piperacillin 100 Tetracycline 30 Tobramycin 10 Trimethoprim 5 Trimethoprim 25 sulfamethoxazole

20-26 27-35 22-28 29-37 19-26 22-30 24-30 22-30 19-27 18-22 17-28 18-24 26-37 19-28 19-29 19-26 24-32

E.coli (ATCC 25922)

P. aeruginosa (ATCC 27853)

19-26 16-22 29-35 15-21 21-27 30-40 19-26 22-28 20-25 28-35 24-30 18-25 18-26 21-28 24-32

18-26 17-23 25-33 16-21 25-33 19-25 -

Quality Assurance in Susceptibility Test The precision and accuracy of the test are controlled by the parallel use of a set of control strains, with known susceptibility to antimicrobial agents. These quality control strains are tested using exactly the same procedure as for the test organisms. The zone sizes shown by the control organisms should fall within the range of diameters given in Table 37.3. When the results regularly fall outside this range, they should be regarded as evidence that a technical error has been introduced into the test, or that the reagents are at fault. Each reagent and each step in the test should then be investigated until the cause of the error has been found and eliminated.

Antimicrobial Susceptibility Testing 145 The quality assurance programme should use standard reference strains of bacteria that are tested in parallel with the clinical culture. They should preferably be run every week, or with every fifth batch of tests, and in addition, every time that a new batch of Mueller-Hinton agar or a new batch of discs is used. The standard strains are: Staphylococcus aureus (ATCC 25923) Escherichia coli (ATCC 25922) Pseudomonas aeruginosa (ATCC 27853) Culture for day-to-day use should be grown on slants of nutrient agar (tryptic soya agar is convenient) and stored in a refrigerator. These should be subcultured onto fresh slants every two weeks. Quality Assurance in Antibiotic Susceptibility Testing Use antibiotic discs of 6 mm diameter. Use correct content of antimicrobial agent per disc. Stock the supply of antimicrobial discs at 20 oC. Use Mueller-Hinton medium for antibiotic sensitivity determination. Use appropriate control cultures. Use standard methodology for the test. Use coded strains from time to time for internal quality control. Keep the antibiotic discs at room temperature for one hour before use. Incubate the sensitivity plates for 16-18 hours before reporting. Incubate the sensitivity plates at 35oC. Space the antibiotic discs properly to avoid overlapping of inhibition zone. Use inoculum size that produces near confluent growth. Ensure an even contact of the antibiotic disc with the inoculated medium. Measure the zone sizes precisely. Interpret the zone sizes by referring to standard charts.

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Practical Manual of Medical Microbiology Table 37.3: Zone sizes with different antimicrobial agents

Antimicrobial agent

When testing against

Disc content

Zone diameter nearest whole mm Resistant Intermediate Susceptible

Enterobacteriaceae Staphylococci Enterococci Pseudomonas Gram negatives Staphylococci Pseudomonas Other Gram negatives Staphylococci Staphylococci Staphylococci Enterococci Pseudomonas Other gram negatives Pseudomonas Other gram negatives

10 µg 10 µg 10 µg 100 µg 100 µg 5 µg 75 µg 75 µg 1 µg 1 µg 10 units 10 units 100 µg 100 µg 75 µg 75 µg

20

COMBINATIONS

20/10 µg

20

20/10 µg 10/10 µg

18

CEPHEMS Cefamandole Cefazolin Cefotaxime Cefoxitin Ceftazidime Ceftizoxime Ceftriaxone Cefuroxime oral Cefuroxime parentral Cephalothin CARBAPENEMS Imipenem MONOBACTAMS Aztreonam

Contd...

Antimicrobial Susceptibility Testing 147 Contd... GLYCOPEPTIDES Telcoplanin Vancomycin

Enterococci Other gram positives

30 µg 30 µg 30 µg

12

30 µg 10 µg

15

30 µg 30 µg 10 µg 10 µg

15

15 µg 15 µg 15 µg

23

30 µg 30 µg 30 µg

19

5 µg 10 µg 10 µg 30 µg 10 µg 5 µg

16

30 µg 2 µg 300 µg 5 µg 250/300 µg 5 µg 1.25/ 23.75 µg

16

AMINOGLYCOSIDES Amikacin Gentamicin

except high resistant enterococci

Kanamycin Netilmycin Streptomycin Tobramycin MACROLIDES Azithromycin Clarithromycin Erythromycin TETRACYCLINES Doxycycline Minocycline Tetracycline QUINOLONES Ciprofloxacin Enoxacin Lomefloxacin Nalidixic acid Norfloxacin Ofloxacin OTHERS Chloramphenicol Clindamycin Nitrofurantoin Rifampin Sulfonamides Trimethoprim Trimethoprim/ sulfamethoxazole

Note: For Vibrio cholerae, the results of disc diffusion tests for ampicillin, tetracycline and trimethoprim/sulfamethoxazole correlate with results obtained by broth microdilution methods.

38 Brucella Agglutination Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE It is an agglutination test for, detection of agglutinins (Brucella) in patient’s serum. Agglutinins detected are usually either lgM or lgG. Materials Required • Antigen: It is available from Izat Nagar or from CPHL, London or it can be prepared from a Brucella abortus strain procured from Institute of Veterinary and Preventive Medicine, Ranipet, • 0.4% Phenol saline. Procedure • Patients serum is diluted 1:20 and then two fold dilutions are done till 1:640 in Phenol saline. • Known positive and negative sera are also diluted in same fashion. • Add 0.5 ml of antigen to each tube of diluted serum and incubate at 37oC for 48 hours.

Brucella Agglutination Test 149 • Appropriate antigen control is also taken. • Results are read by gentle agitation of the deposit and if the supernatant is clear then it is taken as positive. • The end point or titre is the highest dilution of serum causing agglutination. Notes • The agglutination is recorded as 4+. when the agglutination is 100%, 3+when it is 75%, 2+ when it is 50% and 1+ when it is 25%. • Titre is calculated in International Units per millilitre of patient’s serum. For expressing results in IU, double the reciprocal of serum dilution showing 2+ agglutination. If the end point dilution is 1:40 then titre is 80 IU. • Significant titre is 100 IU or above. • H agglutinins tend to persist longer than O agglutinins. Person immunised with TAB vaccine may show high titres of antibodies to all the antigens and so only a marked rise in titre is considered significant.

39 Anti-Streptolysin O (ASO) Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PRINCIPLE It is a neutralisation test used measure the ability of a patient’s serum to neutralise the erythrocyte lysing capability of streptolysin O, an extracellular enzyme produced by Streptococcus pyogenes during infection. Requirements • Antigen: Streptolysin O is available commercially. It is reconstituted before use. • ASO buffer: Phosphate buffered saline. • 5% suspension of RBCs in ASO buffer: Human blood group ‘O’ or rabbit erythrocytes. • Water bath. Procedure • Serum dilution: In a master dilution tube serum is diluted 1:100 by adding 0.05 ml of serum to 4.95 ml of ASO buffer. Subsequent dilutions are made by removing 1 ml of diluted serum from master

Anti-Streptolysin O (ASO) Test 151

• • • • •

dilution tube and adding to another tube in a rack. Then again add 1 ml of ASO buffer in master dilution tube mix and remove 1 ml of this add to the second tube in the rack. Similarly keep on adding 1 ml of buffer each time in master dilution tube and make rest of the dilutions. The dilutions in different serum tubes will be 1:100, 125, 195, 244, 305, 381, 476 and 596. Then add 0.5 ml of the antigen to all the tubes and incubate at 37oC for 15 minutes. After incubation add 0.5 ml of washed 5% RBCs, shake and incubate at 37oC for 1 hour. Controls used with this test are cell control, antigen control and a standard serum control. Readings can be taken as such but centrifugation of tubes makes reading easier. The litre of ASO is the highest serum dilution causing no haemolysis significant titre is considered to be 244 or more.

Notes • Serum should be inactivated at 56oC for 30 minutes before Use • Rise in ASO is detectable after 1 week of infection and very high titres are observed between third and fifth week. • False positive results are due to increase levels of lipoproteins in serum or Contamination of serum by Staph. aureus or Pseudomonas species or by oxidation of ASO. • Commercial kits available for this are Ortho ASO and Rapitex ASL. They are latex agglutination tests.

40 CRP Screen Latex Agglutination Slide Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION C-reactive protein (CRP) is a globulin, which has been so named because of its ability to bind the C-polysaccharide of the cell wall of Streptococcus pneumoniae. CRP is the most responsive of the acute phase proteins, with increased levels usually appearing 6 to 10 hours after injury or infection. Increases in CRP can be much greater than the other proteins, levels over 100 mg/l frequently being found. Thus CRP is a sensitive marker of inflammation and is a useful indicator of the extend of activity in such disorders as rheumatoid arthritis and systemic lupus erythematosus. It is also useful in differentiating bacterial infections from viral infection as CRP levels are elevated only in the former. Principle CRP Screen is based on the principle of latex agglutination. The CRP Latex reagent contains uniform sized latex particles coated with Anti-human C-Reactive protein.

CRP Screen Latex Agglutination Slide Test 153 When a drop of CRP Latex Reagent is mixed with a drop of serum specimen, the latex particles agglutinate to give clear agglutination visible to the naked eye if the concentration of CRP in the serum specimen is more than or equal to 6 mg/l. Requirements 1. 2. 3. 4. 5. 6. 7.

Reagent-1 (CRP Latex reagent) Reagent-2 (Positive control serum) Reagent-3 (Negative control serum) Glass slide Disposable mixing sticks Disposable plastic droppers Rubber teat

SPECIMEN Serum is to be used as the specimen. Fresh specimen should preferebly be used. If necessary, the specimen may be preserved at 2o-8oC upto 48 hours. Discard contaminated or hemolysed sera. No inactivation of the specimen is required. TEST PROCEDURE Bring all the regents to room temperature and mix gently before use. Qualitative Estimation 1. Place one drop of the undiluted serum specimen in one circle on the glass slide with the help of a disposable plastic dropper. 2. Add one drop of Reagent-1 (CRP Latex Reagent) in the same circle on the slide. 3. Mix the undiluted serum specimen with the latex reagent with help of a mixing stick and spread the fluid over the entire area of the circle. 4. Tilt the slide back and forth slowly for 3 minutes and watch for agglutination of latex particles.

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Semiquantitative Estimation Prepare serial two-fold dilutions in physiological saline and repeat steps 1 to 4 as described above. Observe for the presence or absence of agglutination. INTERPRETATION OF RESULTS Qualitative Estimation Clear agglutination of latex particles within 3 minutes is to be interpreted as a positive result, i.e. the CRP titre in the undiluted serum is 6 mg/l or more. No agglutination of latex particles within 3 minutes is to be interpreted as a negative result, i.e. the CRP titre in the undiluted serum is less than 6 mg/l. Semiquantitative Estimation The approximate CRP level in serum sample can be calculated by the following formula: CRP Titre IU/ml = highest dilution with positive reaction × reagent sensitivity (6 mg/ml). QUALITY CONTROL The positive and negative controls provided in the kit need not be run with each specimen. The controls are provided for occasional check of the latex reagent and to differentiate natural granulation of latex reagent from agglutination pattern observed with a serum specimen having elevated levels of CRP. When the test is performed with the positive control, clear agglutination of latex particles is observed within 3 minutes. When the test is performed with the negative control, no agglutination of latex particles is observed within 3 minutes. NOTES 1. Nonspecific positive result may be observed if markedly lipemic, hemolytic or contaminated serum specimen is used. Use of plasma may also give nonspecific positive results.

CRP Screen Latex Agglutination Slide Test 155 2. Slight granularity of latex particles observed occasionally should not be misinterpreted as a positive test result. 3. All reagents of human origin were found negative when tested for HBsAg and HIV anitbody. Handle as if capable of transmitting infection. 4. Avoid contact of the reagents with skin and mucous membrane as these contain sodium azide as preservative. 5. Occasional agglutinations produced after 4 minutes have no diagnostic significance. 6. Rheumatoid factors in the sample may also agglutinate the latex reagent. 7. A prozone phenomenon appears at 80 ug/ml.

41 VDRL Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Veneral disease research Laboratory in USA developed this slide flocculation test for the lab diagnosis of Syphilis. This is a nonspecific standard test for syphilis using non-specific, non-treponemal antigen- cardiolipin, for the detection of antibodies. It is a simple, rapid convenient and economical procedure for seriological testing for syphilis. It has high sensitivity and specificity and can be used for rapid and exact quantitative titration of reactive serum samples. It is well suited for mass serologic surveys and for the examination of large number of serum samples in STD clinics. Nowadays of modification of VDRL test known as RPR (Rapid Plasma reagin) test is used in laboratories for rapid diagnosis. VDRL antigen can be obtained from Laboratories of Serologists, Calcutta, a Government of India Organisation. Requirments 1. VDRL antigen. 2. Glass slide 2 × 3” with 12 paraffin rings approxiamtely 14 mm in diameter (Similiar slides with permanently fixed ceramic rings are commercially available).

VDRL Test 157 3. Buffered Saline 4. Pipettes 5. Microscope • VDRL Antigen is an alcoholic solution containing cardiolopin 0.03%, purified lecithin 0.21% and cholestrol 0.9%. • Buffered saline contains formaldehyde 0.5 ml, Na2H PO4 - 3.037 gm. - 0.170 gm. KH2PO4 NaCl - 10 gm. Distilled water - 1 litre. Preparation of Antigen Pipette 0.4 ml of buffered saline to the bottom of a 10% reagent bottle with flat or concave inner bottom surface. Add 0.5 ml of antigen, drawn from an ampoule into 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 but rapidly so that it takes approximately 6 seconds to complete the delivery. Blow the last drops of the antigen and continue rotation of the bottle for 10 more seconds. Then add 4.1 ml of buffered saline from a 5.0 ml pipette. Stopper the bottle and shake it vigorously for approximately 10 seconds. This emulsion is kept for 30 minutes for maturation. It can be used only within 24 hours. This quantity is sufficient for 250 serum test. Preparation of Serum Clear serum obtained from centrifuging whole clotted blood of the patient is heated for 30 minutes at 56oC. This is to inactivate the Complement activity of the serum which may interfere the reaction. Preliminary Testing of Antigen Emulsion Each preparation of antigen emulsion should first be examined by testing known reactive and non-reactive sera. An unsatisfactory antigen emulsion should not be used.

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Qualitative Serum Test • Pipette 0.05 ml heated serum into a ring of a paraffin ringed glass slide. The serum should spread. • Add one drop (1/60 ml) antigen emulsion on the serum. • Rotate the slide for 4 minutes. Reading and Reporting the Results The tests are read immediately after rotation under a microscope with low power objective. The antigen particles are seen as small fusiform needles which remain more or less evenly dispensed in a non-reactive serum and aggregate in to clumps in reactive serum. No clumps Non-reactive Small clumps Weakly reactive Medium or large clumps Reactive Quantitative Serum Test Quantitative test is performed on all negative serum samples and or all samples showing weakly reactive reactions in the qualitative test. Successive two fold dilutions of the serum are made in 0.9% Saline and each dilution is treated as an individual serum and tested as qualitative serum test. The results are reported in terms of the highest dilution which gives a clear reactive reaction. Notes • VDRL antigen can also be used for the VDRL tube flocculation test and for testing CSF. • The antigen emulsion prepared for any day must not be kept and used for subsequent days. • False positiveness is a common difficulty in VDRL test. It may be due to technical errors or biological reactors. • Biological factors which are responsible for biological false positive (BFP) reaction is found in the following conditions. Malaria, Leprosy, Infectious mononucleosis, disseminated lupus erythematosus, Infective Hepatitis, repeated blood loss, pregnancy, DPT immunisation in children.

42 Treponema Pallidum Haemagglutination Assay (TPHA)

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INTRODUCTION Syphilis is a complex disease which is normally sexually transmitted. The causative organism, Treponema pallidum cannot be cultivated in the laboratory. Infection is normally diagnosed by detecting antibodies specific for T.pallidum in the patient blood or CSF. Two Groups of antibodies are formed. 1. Antibody reacting with non-teponemal antigens used in the VDRL/RPR Tests. 2. Antibody reacting with the specific antigens of Treponema pallidum TPHA is a specific, sensitive passive haemogglutination test for the detection of specific antibodies to Treponema pallidum. PRINCIPLE When diluted positive sample are mixed with sensitised erythrocytes, antibody to the sensitising antigen causes agglutination of the cells.

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The cells using a microtitration diluter mix row-l and transfer 1 volume to row-2 mix and transfer 1 volume to row-3. Mix and discard 1 volume from row-3. Transfer a second volume from row-3 to row4, mix and discard 1 volume from row-4. • Add one drop (75 microlit) of test cells to row-4 using the attached cell dropped and one drop (75 microlit) of control cells to row-3 using the cell dropper. Tap plate gently to mix. This results in final serum dilutions in row 3 and 4 1/40 to 1/80 (Fig. 42.1).

Figure 42.1: Serological dilution tray

• Cover and let stand for 45-60 minutes. • Examine for agglutination patterns. Quantitative Test Quantitative tests may be modified by omitting the control cells and preparing only one final dilution. The titre is the highest dilution showing agglutination. Observation and Results • Agglutinated cells form an even layer over the bottom of the well. Non-agglutinated cells form a compact button in the centre of the well. Weak agglutinated cells form a characteristic ring pattern.

Treponema Pallidum Haemagglutination Assay (TPHA) 161 • Agglutination of the test cells but not the control cells indicates the presence of specific antibody to T.Pallidum. Observation

Results

Agglutination of test cells and non-agglutination of control cells

Reactive

No agglutination of test cells and control cells

Non-reactive

43 Widal Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Enteric fever in man is caused by Salmonella typhi (Typhoid) and Salmonella paratyphi (Paratyphoid). S. paratyphi is sub divided in to 3 sero types, S. paratyphi A, S. paratyphi B and S. paratyphi C. S. paratyphi. C is very rare in India. The lab diagnosis of enteric fever includes Blood culture, Stool Culture and Serological test. The common serological test used is known as Widal Test. It is tube agglutination test. PRINCIPLE It is the test for detection of antibodies produced by the host against antigenic determinants on the surface of the bacterial agents in response to infection with Salmonella typhi and S. paratyphi. These antibodies are agglutinating antibodies as they result in agglutination of the bacterial antigens. The patient serum is tested by agglutination for its titres of antibodies against H and ‘O’antigen suspensions of S. typhi and S. paratyphi. Requirement • Small test tubes.

Widal Test 163 • Antigen suspensions: These may be prepared from suitable stock cultures in the laboratory but generally commercially prepared suspensions are used. • 0.85% NaCl. • Water bath. Procedure (Method) • Patient’s serum is tested in a series of dilution against each of the different antigen suspensions. • Take 7 tubes for each series. Use 6 for serum dilution and 7th for nonserum control. • Add 0.4 ml of Saline in tubes 2 to 7. Dilute patient serum 1 in 15 in Saline and add 0.4 ml of these to tubes of tubes 1 and 2. Dilution of Serum in tube 2 will be 1 in 30 (vol 0.8 ml) • Mix the contents of the tube 2 thoroughly and transfer 0.4 ml to tube 3. Repeat the process till tube 6 and discard 0.4 ml in the end. The serum dilution in 6 tubes will be 1:15, 1:30, 1:60, 1:120, 1:240 and 1:480 and seventh tubes serves as a negative control. • Add 0.4 ml of antigen suspension to each tube starts from tube 7 and working backwards to tube 1 so, the serum dilution in tube 1 to 6 will be now 1.30 – 1.960 the four different antigens used are TO, TH, AH and BH . • Incubate the tubes in a rack in water bath. The tubes with somatic ‘O’ antigen incubater at 50 oC for 2 hours or 37 oC for 4 hours. Observation and Result • The results are read by viewing the tubes under good light against a dark backround with the aid of a magnifying lens. In case of ‘H’ agglutinins large flakes are formed and with ‘O’ agglutinins only small granules are formed. It necessary, the tube can be gently rotated, to swirl up granules from deposit. • The titre of the serum is the highest dilution of serum giving visible agglutination.

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Notes • A Progressive rise in the titre between 1st and 3rd week after onset of fever is highly significant. A positive or negative result in a single test is not significant. • Since the antibodies are detected only after 7 to 10 days of illness, test should be done later. • The serum of some uninfected subjects cause agglutination at dilution of about 1 in 50. So, titeres are considered significant when agglutination occurs in serum dilution above 100. • H agglutinins tend to persist longer than 0 agglutinins. Persons immunised with TAB Vaccine may show high titres of antibodies to all the antigens and so only a marked rise in the titre is considered significant.

44 Enzyme Linked Immunosorbent Assay (ELISA)

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ELISA is a heterogenous enzyme assay that uses solid phase, e.g. plastic tubes, polyvinyl chloride plates, beads, microplates, membranes such as cellulose acetate or cellulose nitrate. A range of enzyme labels and substrates for each enzyme are available. The most popular enzymes are horse radish perioxidase (HRP) and alkaline phosphatase (AP) and substrates are orthophenylene diamine (OPD) or diaminobenzidine (DAB) or orthotoludiene (OTD) and p-Bromochloro indoly phosphate-(BCIP) respectively. Occassionally B-galactosidase enzyme has also been used, and the substrate is o-nitrophenyl galactoside (or Bromochloroindolyl-DGalactopyranoside (BCIG). ELISA FOR ANTIBODY DETECTION The indirect method is most commonly used for antibody detection. Antigen is first absorbed on to a solid phase, most conveniently a

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microtiter plate, then incubated with test serum and washed to remove any unbound components. The amount of antibody bound is measured by addition of enzyme-linked anti-human immunoglobulin (class, subclass specific) and an appropriate substrate to produce a coloured end product. The depth of colour is proportional to the amount of specific antibody in the serum. Since, the best all round system is the combination of horseradish peroxidase with the substrate H2O2 and the indicator OPD as it is inexpensive and extremely sensitive this will be discussed further. Reagents Carbonate buffer (pH-9.6) Solution A : 5.3 gm Na2CO3 dissolved in one litre of double distilled water. Solution B : 4.2 gm NaHCO3 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. Washing buffer (PBS-Tween-20) Phosphate buffered saline (0.01M) (PBS) pH 7.2 with 0.05% Tween 20. Conjugate Goat antimouse immunoglobulin conjugatedwith HRP (Dakopatts, Denmark). Citric acid phospate buffer (0.1M) pH-5.0 Citric acid : 7.3 gm Na2HPO4 : 11.86 gm Dissolve in litre double distilled water and adjust pH to 5.0. Substrate Orthophyenylene diamine: 8 mg (Sigma chemicals, Co, USA). Citrate buffer: 15 ml Hydrogen perioxide: 15 ml 4% Bovine serum albumin (BSA) in PBS – Tween – 20. 3NH2SO4

Enzyme Linked Immunosorbent Assay (ELISA) 167 Procedure • Make up antigen to optimal dilution in carbonate buffer and add 100 μl per well in 96 well microtilter plate and incubate at 37oC for 3 hours or preferably at 4oC overnight. • Decant the antigen solution and wash the plates thrice with washing buffer. • Remove all residual fluid with rigorous shaking. • Add 100 μl of 4% BSA in washing buffer to each well and incubate at 37oC for one hour. • Wash thrice with washing buffer. • Add 100 μl of diluted serum samples and incubate for one hour at 37oC. • Repeat the washing procedure. • Add 100 μl of conjugate (diluted 1:500 or 1:1000 or as recommended by the manufacturers) to each well and incubate as before. • After three washings, add 100 μl of substrate solution and incubate for another 30 minutes at 37oC. This results in development of yellow colour. • Stop the reaction with 50 μl of 3N H2SO4 to each well. • Read the results with naked eye or at 492 nm in an ELISA reader. Notes • Preliminary chequer board titrations are carried out for standardising antigen and antibody concentration, where antigen in increasing protein concentration (5, 10, 20, 30, 40 and 50 μg/ ml) in titrated against serial dilutions of known positive and negative sera. • Do not add azide to PBS or sera as its ions inhibit peroxidase reaction. • Read all plates at a standard time after addition of a substrate (e.g. 30 minutes) because the colour is not very stable. • The substrate should be made just before use as it decomposes spontaneously is unstable, and darkens with exposure to light. • For plate based assays, use micropipettes (Figs 44.1 and 44.2).

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Figure 44.1: Micro tips

Figure 44.2: Micropipette

Enzyme Linked Immunosorbent Assay (ELISA) 169 • OPD should be stored at 4oC in the dark. It is carcinogenic, so gloves should be worn while using it. • Appropriate control, such as controls with positive and negative sera, buffer control, conjugate control and substrate control should be tested along with every plate. Elisa for Antigen Detection This involves antibody coated plates to immobilise target antigen, which is then detected with a second antibody directed to a second site on the same antigen. The second antibody is generally directly enzyme labelled as anti-immunoglobulin conjugates cannot be used when both “capture” and “indicator” antibodies are from the same animal species. Reagents • • • • • • • • •

Carbonate buffer Blocking solution Washing buffer Monocional anitbodies (MAb) For coating plates: 1-10 μg/ml in carbonate buffer (or as determined to be optimal). Prepare 10 ml for each plate. For conjugated MAb: the dilution must be determined by titration. Typical conjugates work at 1/2000 dilution. Substrate solution.

Procedure • Coat the ELISA plates with MAb in carbonate buffer by adding 100 μl to each well, and incubate overnight at 4oC. • For each ELISA Plate, set up a replica plate containing serum samples and enzyme conjugated monoclonal. The contents will later be transferred to the ELISA plate. To each well mix 50 μl of diluted serum and 50 μl of diluted conjugate, each at 2X desired final concentration Incubate overnight at 4oC.

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• Remove MAb from ELISA plate and add 100 μl of blocking solution. Incubate for 30 minutes at 37oC and then wash three times in washing buffer. • Transfer contents of replica plate into corresponding well of ELISA plate. • Incubate for 6 hour at 37oC, after sealing plate. • Wash plates three times in washing buffer. • Add 100 μl of substrate solution in each well. • Incubate at 37oC for 30 minutes and stop the reaction with 50 μl of 3NH2SO4. Take the readings at 492 nm in ELISA reader. Note This procedure, in which the circulating antigen and the conjugated monoclonals are first co-incubated, gives a greater sensitivity with serum samples, perhaps because the conditions favour dissociation of immune complexes and binding of enzyme linked monoclonal. However, if assaying free antigen, then a simpler sequential protocol can be adopted in which the coated ELISA plate is first incubated with test antigen, then washed and exposed to conjugated monoclonal, washed again and assay with substrate solution. TYPES OF ELISA Stick ELISA (Indirect) This is a modification of indirect method for antibody detection. In this, the antigen is immobilised on cellulose acetate membrane square fixed onto a plastic stick. The test screen is then incubated and the antibody bound to immobilised antigen is detected with enzyme labelled second antibody. The result is revealed by the development or disappearance of colour using a suitable substrate. Stick ELISA (Sandwich) In the sandwich ELISA, used for antigen detection immunoglobulin containing specific antibody is immobilised on CAM Squares. The test sample is then incubated and the bound antigen is detected

Enzyme Linked Immunosorbent Assay (ELISA) 171 using the enzyme conjugate of same antibody used for sensitisation of CAM squares. Competitive (ELISA) This is useful for identification and quantitation of either antigen or antibody. In antigen determination by this method, antigen present in the sample competes for sites on the antibody with labelled antigen added to the medium. The colour change will be inversely proportional to the amount of antigen in the sample. Inhibition ELISA This works similiar to that of competitive ELISA, but in this system the two antigens (antigen in test sample and enzyme labelled antigen) are added one after another. This is useful especially when test serum contains both antigen and antibody of interest in immune reaction. Inhibition ELISA is also useful in determining the identity of antigen or antibody. Note Stick ELISA is simpler and economic as ELISA plates are not required which are imported and costly.

45 Experimental Animals ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

The animal inoculation is an important part in determining the pathogenicity of virulence of the organisms. The commonest animals used for experimental purpose in microbiology are: 1. Rabbits 2. Guinea pigs 3. Rats 4. Mice Rabbits (Fig. 45.1) 1. Used for exaltation of neurotropic virus by subdural injection, e.g. Rabies virus. 2. Maintenance of Nichols strain of Treponema pallidum by intra testicular injection. 3. To differentiate between the human and bovine strains of Tubercle bacilli Bovine strain : generalized lesion. Human strain : localized lesion. 4. Preparation of amboceptor for serological tests. 5. Collection of blood for media preparation by cardiac puncture.

Experimental Animals 173

Figure 45.1: Rabbit

The modes of inoculations are subcutaneous, intravenous intraperitoneal, intracerebral and intratesticular. The marginal ear vein is selected for intravenous inoculation. Guinea Pigs (Fig. 45.2) Used for 1. Diagnosis of diseases like tuberculosis, diphtheria. 2. Collection of complement from its blood. Modes of inoculation are intraperitoneal and intramuscular.

Figure 45.2: Guinea pig

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Rats (Fig. 45.3) 1. Study of murine typhus and plague. 2. For endocrinological assays, e.g.: Oral contraceptives. Modes of inoculation are by intravenous (Tail vein), subcutaneous and intraperitoneal routes

Figure 45.3: Rat

Mice (Fig. 45.4) 1. For pure isolation of D. pneumoniae. 2. For demonstration of capsule sweeling reaction and capsule can be demonstrated in tissues. 3. For immune tolerance reaction. 4. For experimental cancer research.

Figure 45.4: Mice cage

Figure 45.5: Mouse

46 Potassium Hydroxide Wet Mount ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Potassium hydroxide (KOH) wet mount preparation is very useful for the laboratory diagnosis of fungal infection. The KOH clears out the background scales or cell membranes that may be confused with fungal hyphal elements in the microscopy of clinical specimens like hair, nail or skin. Warming over a low flame hastens the digestion of the keratin. Requirements 1. 2. 3. 4. 5. 6. 7.

10% KOH Coverslip Glass slide Needle Bent wire Bunsen flame Specimen

Method Two types methods are used a. Slide KOH mount b. Tube KOH

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Slide KOH Mount • Place the specimens like epidermal scales, nail, hair, skin scrapping or tissue on a cleanslide. • Pour a drop of 10% KOH on the specimen and emulsify • Place a converslip over it. • Heat it gently over a flame • Leave the slide for 5-10 minutes. • Examine the slide under microscope tube KOH. It is mainly done for the biopsy specimens in 10% KOH the homogenesed tissue material is dissolved and examined after keeping for an overnight at room temperature. Observation Shining fungal elements can be observed under microscope. Different Morphological forms of fungi can be clearly seen in a KOH wet mount. Identification of fungi is done by the characteristic morphology.

47 Lactophenol Cotton Blue Mount ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Wet mount preparations using lactophenol cotton blue (LPCB) are very useful to study the morphology of filamentous fungi. LPCB–ingredients and Preparation Phenol 20 ml Lactic acid 20 ml Glycerol 40 ml Distilled water 20 ml Mix the reagents thoroughly. Dissolve 0.05 gm of cotton blue stain in distilled water and add to the reagents. The Phenol acts as disinfectant, Lactic acid preserve the morphology of the fungi and glycerol is hygroscopic agent which prevents drying. Requirements 1. Fungal culture 2. Teasing needles 3. Coverglass

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Figure 47.1: LPCB mount of Pencillium

4. Glass slide 5. Nail polish Method • Place a drop of LPCB on a clean glass slide. • With the help of an ‘L’ shaped wire pick up a small portion of the fungal colony and place it in the drop of LPCB. • With teasing needle, tease the fungal culture and spread in the LPCB. • Place a coverslip over the stain and observe under 10 × or 40 × objective. • Permanent LPCB mount of the specimen can be made by sealing the edge of the coverslip with nail polish (Fig. 47.1).

48 Culture of Fungi ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION The isolation of Fungi is not difficult. The essential requirement is a source of nitrogen in the isolation medium. The medium commonly used is Sabouraud’s dextrose agar. Broth media are not recommended except for blood culture. Many fungal pathogens have optimum growth temperature below 37oC. Two culture media are inoculated and incubated at different temperatures. This procedure is useful to identify dimorphic fungi. Mycelial Phase develops at 25oC and yeast phase develops at 37oC. Fungi grow relatively slow and cultures should be retained for at least 4 weeks. The media may be prepared in test tube slopes or Petri dishes. The slopes are preferred as compared to plates for cultivation of Fungi as they are safer, require less space and are more resistant to dehydration during prolonged incubation. Culture Media The following culture media are commonly used in the mycology laboratory.

180 1. 2. 3. 4. 5. 6. 7. 8. 9. 10.

Practical Manual of Medical Microbiology Sabouraud’s dextrose agar Neutral Sabouraud’s dextrose agar Sabouraud’s dextrose agar with antibiotics. Corn meal agar Rice starch agar Brain heart infusion agar Bird seed agar Czapek’s dox agar Malt peptone agar. Potato dextrose agar.

Sabouraud’s Dextrose agar Dextrose 40 gm Peptone 10 gm Agar 20 gm Distilled water 1litre Adjust pH to 5.5 autoclave at 121oC for 15 min. Dispense the medium in sterile petridishes or test tubes. Overgrowth of saprophytic fungi and bacteria make the identification of pathogen difficult. Sabouraud’s Dextrose Agar with Antibiotics To the SDA while boiling add the following antibiotics. Cycloheximide 500 mg Chloramphenicol 50 mg Gentamicin 20 mg. Dissolve cycloheximide in 10 ml acetone; add to the boiling medium and mix. Dissolve chloramphenicol/Gentamicin in 10 ml of 95% alcohol and add to the boiling medium. Remove from heating and mix well. Dispense in tubes and autoclave at 121oC for 15 minutes. Gentamicin and Chloramphenicol prevent the bacterial contamination and cycloheximide inhibit saprophyte Fungi. Cycloheximide should not be used to isolate Cryptococcus and Aspergillus because these fungi are sensitive to this antibiotic (Fig. 48.1).

Culture of Fungi 181

Figure 48.1: SDA plate with Candida colonies

49 Fungal Slide Culture (Riddle's Method) ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Slide culture is used to study the undisturbed morphological details of the fungi. The difficulties faced during direct observation of fungal colony are overcome by this method. Teasing and separating fungal colony may break the reproductive structures from mycelia and this will pose a problem in the identification of Fungus. Requirements (Fig. 49.1) 1. 2. 3. 4. 5. 6. 7. 8. 9.

Glass slide Bent glass rod (V shaped) Sterile Sabouraud’s dextrose agar prepared in a petridish. Knife Coverslip Filter paper Water BOD incubator Fungal culture

Fungal Slide Culture (Riddle's Method) 183

Figure 49.1: Slide culture set

Method • Sterilise slide culture test set by hot air oven (slide cuture set consist of a petridish, inside the petridish keep a filter paper, place the ‘v’ rod . on V rod place a glass slide and converslip. • Cut out one square cm block of SDA from the sterile petritish using a sterile knife • Place the agar block in the centre of the slide in the slide culture set. • With bend wire or straight wire pick up some fragments of fungal colony from the culture. • Inoculate the fungal fragments around the periphery of the agar block. • Place a coverslip on the agar block with the help of sterile forceps • Moisten the filter paper with sterile water. • Close the petridish and incubate at room temperature (BOD incubator) – minimum 48 hours. • When growth appears beneath the coverslip, take a slide, place one drop of LPCB on it,and place the coverslip removed from the block on the LPCB. • Observe under microscope. Observation Fungal hyphae and reproductive structures can be easily identified.

50 Identificationof Fungal Isolates ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Unlike bacteria the biochemical tests are not useful or not available for identification Fungi. Fungi are mainly identified by their morphological features. Based on Morphology Fungi are classified into four groups 1. Yeast, e.g. Cryptococcus 2. Yeast like, e.g. : Candida 3. Filamentous or molds, e.g. : Aspergillus, mucor 4. Dimorphic Fungi, e.g. Histoplasma. Methods 1. Macroscopic examination 2. Microscopic examination 3. Biochemical test. Macroscopic Examination Noting the characteristic growth and colony characters of fungi are

Identification of Fungal Isolates 185 useful for the identification of genus. Note the colour and Texture of the growth it is useful for the presumptive diagnosis. Microscopic Examination LPCB mount is commonly used for the microscopic examination of Fungi. Special staining methods like Gram’s staining, Indian ink Colcofluor white stain, etc. can also be performed in special occasions. Morphological criteria are useful for the identification of Fungi rather than biochemical tests. Note the Characteristics like • • • • • • • • •

Cell shape Cell size Hyphae – septate or non-septate Branching Rhizoids Capsule Pseudomycelium Germtube Asexual and sexual reproductive structures. These characters will be useful in identification of individual fungi. The morphology of and their identification points medically important fungi are given in this chapter. Cryptococcus Neoformans • A soil Saprophyte, abundant in the feces of pigeons and other birds • Cryptococcal meningits and pulmonary Cryptococcosis are seen in immuno compromised patients like HIV infected. • Yeast fungi • Budding yeast cells can be demonstrated in the specimens (Fig. 50.1). • Characterised by presence of polysaccharide capsule. This can be demonstrated by Indian ink preparation. • On SDA fungus grows and forms smooth, mucoid, cream coloured colonies.

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Figure 50.1: Cryptococcus

• Hydrolysis of urea (urease test) differentiates C.neoformans from Non-pathogenic species and Candida. • Gram’s Stained smear of culture will show gram positive, round or ovoid budding cells having 4-20 μm in diameter. Candida Albicans (Figs 50.2 to 50.5) • • • • • • • • •

Yeast like fungi Normal inhabitants of the skin and mucosa. Candidiasis is an opportunistic endogenous infection. Gram positive budding yeast cells can be demonstrated (Fig. 50.3). On SDA colonies are creamy white, smooth and with a yeasty odour. They form chlamydospores on corn meal agar cultures at 20oC (Fig.50.4) C.albicans forms germ tube. Pseudohyphae can also be demonstrated in cultures. Do not hydrolyse urea.

Identification of Fungal Isolates 187

Figure 50.2: Candida

Figure 50.3: Candida in a Gram’s smear

Figure 50.4: Chlamydospores of Candida

Trichophyton rubrum • Trichophyton is a dermatophyte • They infect skin, hair and nails and produce tinea or ring worm infection. • Colonies on SDA are powdery, Velvety or Waxy. produce red pigment. • Septate, branching filamentous hyphae (Figs 50.5 to 50.7) • Two types of asexual spores are produced, microconidia and macroconidia.

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Figure 50.5: Conidia of dermatophytes

Figure 50.6: Trichophyton species

Identification of Fungal Isolates 189

Figure 50.7: Trichophyton species

• Microconidia are abundant. They are arranged in clusters along the hyphae or borne on conidiosphores. • Macroconidia are relatively scanty. They are elongated with blunt ends. Pencil shaped macroconidia differentiates this genus from other dermatophytes. Microsporum canis • • • •

Dermatophyte fungi They infect hair and skin not nails and produces Tinea. Colonies are cottony, Velvety or powdery with orange pigment. Septate branching filamentous hyphae

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• Microconidia are scanty • Numerous thick walled 8 to 15 celled Macroconidia are the Predominant spore form (Fig. 50.8). • Spindle shaped macroconidia (Fuse form) are borne singly on the ends of hyphae.

Figure 50.8: Microsporum species

Epidermophyton floccosum • • • •

Dermatophyte fungi They infect skin and nail not the hair to produce Tinea. Colonies are powdery and greenish yellow in colour. Microconidia are absent.

Identification of Fungal Isolates 191 • Macrocondia are multicellular, club or pear shaped and arranged inclusters. • Septate branching filamentous hyphae. • Numerous thick walled 8 to 15 celled Macroconidia are the predominant spore form (Fig. 50.9).

Figure 50.9: Epidermophyton

Aspergillus • Saprophyte fungi produce opportunistic infections. • The commonest human disease is otomycosis. It can also produce pulmonary aspergillosis and disseminated aspergillosis • They are common lab contaminants of culture media • Sepatate branching hyphae • Asexual conidia are arranged in chains. Conidia are formed on elongated cells called sterigmata. Sterigmata are borne on the expanded ends of Conidiophores. • The species indentification of Aspergillus is done by the morphology and growth characters (Figs 50.10 and 50.11). • The colonies are velvety, yellow to green or brown. • Important species are A. flavus A. fumigatus A. niger

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Figure 50.10: Aspergillus flavus

Figure 50.11: Aspergillus species

Identification of Fungal Isolates 193 Pencillium • Saprophytic fungi • Lab contaminant • Pencillium marneffei is the only pathogen reported which causes opportunistic systemic mycosis in AIDS Patients. • Septate branching hyphae • Conidophores arise in various forms producing phialides giving a brush like arrangement (Fig. 50.12). • The Conidia are unicellular and in chains with youngest at the base. • Sometimes Conidiophores are aggregated into stalks (Sterigmata) which are called coremia.

Figure 50.12: Pencillium

Rhizopus • Common Bread Mold • Produces opportunistic systemic fungal infection like mucoromycosis. • The colonies are dense and have a hairy appearance on sporulation the growth turns to black. • Rhizopus has non-septate hyphae. • Branching of hyphae is noted. • From the aerial hyphae root like projections called Rhizoids arise to the medium (Fig. 50.13).

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Figure 50.13: Rhizopus

• Unbranched sporangiophores arise opposite to rhizoids • Important species are R. arrhizus R. microsporus. Mucor • Saprophytic fungi related to Rhizopus (order mucorales) • Produce opportunities systemic mycosis like Rhinocerebro mucoromycosis. • Non-septate branched hyphae • The colonies are dense and have a hairy appearnce. • Sporangiophores are branched. They terminate in large globose sporangium containing numerous spores Rhizoids are absent (Fig.50.14).

Figure 50.14: Mucor

51 Germ Tube Test ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Candidiasis is the commonest fungal disease affecting mucosa, skin and its appendages and the internal organs. Candidiasis is caused by various species of yeast like fungi belonging to the genus candida. Candida species are normal commensals of the human beings. They produce opportunities infections especially in immunocompromised patients. There are more than 200 specis of Candida. The commonest pathogenic species of this genus is C. albicans which is responsible for about 90% of candidal infection. Germtube is a long tube like projection extending from the yeast cell. There is no constriction at the point of attachment to the yeast cell. Germ tubes are formed only by Candida albicans with in two hours of incubation in sheep or normal human serum incubaled at 37oC. Demonstration of germ tube is also known as Reynolds-Braude Phenomenon (Fig. 51.1). Germtube test is commonly used to identify C. albicans in the laboratory. Requirements 1. 24 hours culture of suspected Candida colony on SDA (Sabouraud’s Dextrose Agar).

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Figure 51.1: Germ tube formation

2. Human/Sheep serum. 3. Test-tubes. Method • • • • •

Take 0.5 ml of serum in a sterile test tube. Using a sterile wireloop take a colony of fungi to be tested. Mix the colony with serum in the test tube. Incubate the tube at 37oC for 2 hours. After incubation place one drop of suspension from the tube on to a glass slide. • Place coverslip and examine under microscope. Observation Germ tubes are seen as long tube like projections extending from yeastcells. Note Germ tubes are also produced by few strains of Candida stellatoidea and C. tropicalis.

52 Diagnosis of VirusInfections ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

The viruses are usually invisible under ordinary microscopic examination. Measured in Mu or nm (Millimicrons or nanometer). They do not grow on non living culture medium. Laboratory diagnosis of viral infections is of limited use in the case of the individual patient but it’s important in the field of epidemiology and public health as its helps to control the outbreak of the infection. Laboratory support is also required for field trials and subsequent surveillance of vaccines. Diagnosis of virus infections depends on three principle techniques: 1. Direct demonstrations of virus in smear from lesions. 2. Isolation and identification of virus. 3. Serological tests A. Direct demonstration of virus : a. By using electron microscope. b. By flourescent antibody tests. c . By demonstration of virus inclusion bodies. B. Isolation and identification of virus : a. The viruses are isolated by inoculating into (i) Tissue culture (ii) Chick embryo (iii) Laboratory animals. b. The virus that is grown is identified by neutralisation tests, haemadsorption and plaque formation, etc.

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Serology Infection is diagnosed by the demonstration of the development of virus antibody in the patient serum. Usually two samples of blood are collected, the first one taken during the acute phase of the illness and the second taken 10-14 days later in convalescence. The acute and convalescent serum samples are tested in parallel to detect a rise in the titre of antibodies to the virus. Recent infections are indicated by a rise in antibody titre from the acute to the convalescent serum samples. Three main serological tests are used to detect antibodies. 1. Complement fixation tests. 2. Haemagglutination inhibition tests. 3. Neutralization tests. Embryonated Egg Inoculation (Figs 52.1 and 52.2) Egg inoculation and cultivation of viruses is useful for the production of vaccines. The embroynated egg offers several sites for the cultivation of viruses. Inoculation sites available in a ten day old embryonated egg are: 1. Choriallantoic membrane (CAM): Inoculation on the CAM produces visible lesions (POCKS) Different viruses have different pack morphology under optimal conditions, each infections virus can form one. Pock counting can be used for the assay of viruses (Fig. 52.1). 2. Allantoic cavity 3. Amniotic sac 4. Yolk sac Tissue Culture Cell cultures are commonly used in virology lab for cultivation of viruses (Fig. 52.3). The available cell lines for common use are mentioned below. 1. Primary cell cultures: a. Rhesus monkey kidney cell culture b. Human amnion cell culture c . Chick embryo fibroblast cell culture

Diagnosis of Virus Infections 199

Figure 52.1: Embryonated egg and inoculation of CAM

Figure 52.2: Inoculation of embryonated egg–into amniotic cavity, allantoic cavity and Yolk sac

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Figure 52.3: Tissue culture bottle

2. Diploid cell strains a. WI-38 b. HL-8 3. Continuous cell lines a. HeLa b. HEP-2 c . KB d. McCoy e . Detroit-6 . f Vero g. BHK-21

53 Lab Diagnosis of Malaria ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Malaria is a febrile illness caused by the presence of malaria parasites (Plasmodium sps.) in the human body. The parasite is found in the red blood cells of human beings. A person suffering from malaria will always get fever but all fevers are not due to malaria. The diagnosis of malaria is dependent upon the demonstration of malaria parasite in the peripheral blood film. The essential requirements for an accurate diagnosis of a case of malaria are: • Making a proper blood smear from a patient having fever • High quality staining of the smear • Examination of the stained slide by a technician skilled in malaria microscopy. Preparation of Blood Smear Malaria microscopy is the key to the diagnosis of malaria. Requirements Clean glass slides Pricking needle (Hagedorn triangular no. 12) Spirit Cotton

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Slide box Lead pencil Two types of smears are prepared from the peripheral blood – one thin smear and the other thick smear. Thick film examination is about 20 times more sensitive than thin film examination for parasite detection. Thin film examination is done for finding out the species of Plasmodium. Prepare the thin and thick blood smears in the following way (Figs 53.1 to 53.7): Hold the third finger of the left hand of the patient between your left thumb and finger at the first phalangeal joint. Wipe finger tip with swab dipped in savlon solution. Allow the finger tip to dry. Hold the pricking needle (Hagedorn triangular no.12) in right hand and prick the finger and allow blood drop to ooze out. Take a clean, dust free, grease free slide and take 3 drops of the blood 1 cm from the edge of the glass slide. Take another drop of blood 1 cm from the first drop of blood. Take another clean slide with smooth edges and use it as a spreader. Make thick smear by joining the 3 drops of blood and spreading it in an area of 10 mm diameter. Make thin smear by bringing in contact the spreader with the drop of blood at an angle of 30-45o from the horizontal and pushing the spreader steadily down the surface of the slide drawing the blood behind till the smear is formed (Fig. 53.7). Allow it to air dry. Put the slide number on thin smear with lead pencil. Qualities of good thick and thin smears are given below: Thick smear It should be 10 mm away from the edge of the slide. It is round in shape with a diameter of 10 mm. Its thickness contains 10 layers of RBCs. 10-12 WBCs should be visible per oil immersion field of microscope.

Lab Diagnosis of Malaria 203

Figure 53.1

Figure 53.2

Figure 53.3

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Figure 53.4

Figure 53.5

Figure 53.6

Figure 53.7 Figures 53.1 to 53.7: Preparation of peripheral blood smear

Lab Diagnosis of Malaria 205 Thin Smear Uniformly spread over the slide. Thin enough so that newsprint can be read through the smear. It is tongue shaped. Consists of a single layer of RBCs. Also used to label the identity of the patient. Staining of blood films Giemsa stain which is a mixture of eosin and methylene blue is widely used for staining the films. In some countries, JSB staining is used in the National Malaria Programme. Giemsa Staining Use slides which are well dried, peferably overnight. Fix thin film by dabbing it gently with a small piece of cotton dampened with methanol. Avoid methanol coming in contact with the thick film. Place the slides back to back in a staining trough making sure that all thick films are at one end of the trough. Prepare a 3% solution of Giemsa stain by adding 3 ml of Giemsa stock solution to 97 ml of buffered water. Pour the stain gently into the trough until the slides are totally covered. Do not pour the stain directly on to the thick films. Leave the slides in the stain for 30-45 minutes. Pour clean water gently into the trough to float off the scum on the surface of the stain. While pouring water, do not disturb the thick films. Pour off the remaining stain gently and rinse again in clean water for a few seconds. Pour off the water. Remove the slides one by one and place them, film side downwards, in a drying rack to drain and dry, making sure that the thick film does not touch the edge of the rack. Examination of the Stained Slides Since it takes almost 10 times as long to examine a thin film as to examine a thick film, in practice thick film is examined first. Thin

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film is examined only when the thick film gets autofixed or when it is necessary to confirm the identification of a species. Examination of the Thick Film Using the 40 x objective, select a part of the film that is well stained, free of staining debris and is well populated with WBCs. Place a drop of immersion oil on the thick film. Lower the 100 x oil immersion objectives over the selected portion of the blood film, so that it touches the immersion oil. Confirm that the portion of the film is acceptable and examine the slide for 100 oil immersion fields by moving along the width of theslide. Examine atleast 100 good fields before a slide can be pronounced negative. Record your findings in an appropriate form. Examination of Thin Film Place the slide on the mechanical stage and position the 100 x oil immersion objective over the edge of the middle of the film. Place a drop of immersion oil on the edge of the middle of the film. Lower the oil immersion objective until it touches the immersion oil. Examine the blood film by moving along the edge of the thin film, then moving the slide inwards by one field, returning in a lateral movement and so on. Continue the examination, for atleast l00 fields to determine whether the blood film is positive or negative for malaria. Recognition of the Malaria Parasite There are four species of malaria parasite namely Plasmodium vivax (commonest), P. falciparum (second common spp.) P. malariae (less common) and P. ovale (rare). In all stages of development, the parasite will stain the same colour with Giemsa stain. These are: Chromatin which is part of the parasite nucleus is usually round in shape and stains a deep red.

Lab Diagnosis of Malaria 207 Cytoplasm occurs in a number of forms, from a ring shape to a totally irregular shape. It always stains blue, although the shade of the blue may vary between the parasite species. The simplest guide to distinguishing between the four species of malaria is the effect the parasite has on infected red blood cells, whether or not it is enlarged and whether or not staining reveals Schuffner’s dots or Maurer’s dots. There are no RBCs in the stained thick blood film. The malaria parasites can be seen though they appear to be smaller than in thin blood films (Figs 53.8 to 53.10).

Figure 53.8: Morphology of malarial parasites

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The essential morphological features distinguishing the four species of malaria parasite are given in Table 53.1 and Fig. 53.8. Table 53.1: Differential characters of malaria parasites P. falciparum

P. vivax

P. malariae

P.ovale

Size

Not enlarged

Shape

Round and sometimes crenated

Enlarged

Not enlarged

Enlarged

Round or oval (frequently bizarre form)

Round

Round or oval,often fimbriated

Colour

Normal but may become darkened

Normal but inclined to be pale

Normal

Normal

Stippling

Maurer’s dots (large red spots)

Schuffner’s dots (small red dots)

Ziemann’s dots (few tiny dots not important for diagnosis)

James’s dots (numerous small red dots)

Pigment

Usually black or very dark brown

Fine golden brown granules seen in cytoplasm

Black or brown coarse granules

Resemble more closely P. malariae

Small,compact dark, staining parasite. Multiple infections of single RBC

Large light staining parasite. Many trophozoites, may be amoeboid

Regular shape Regular shape. Size and moderate size. in between P. vivax Strong tendency to and P. malariae form a band across the infected RBC

Common. Only rings and Stages found gametocytes in smear

Trophozoites, Schizonts, Gametocytes

As in P. vivax

As in P. vivax

Ring stage

Large 2.5 µm, usually single. Prominent thicker chromatin

Similar to P. vivax but thicker

Similar to P. vivax, more compact

Trophozoite Compact, small, vacuole inconspicous, seldom seen in smear

Large, irregular actively amoeboid prominent vacuole Chromatin as dots or threads

Characteristic band form, vacuole inconspicous

Compact coarse pigment, chromatin as large irregular clumps

Schizont

Small, compact seldom seen in blood smear

Large, filling the RBC, segmented, yellow brown pigment

Nearly fills RBC, like segmented, daisy head, pigment is dark brown

Fills three fourth of RBC, segmented Dark yellow brown pigment

Microgametocyte

Larger than RBC, kidney shaped with blunt round ends, cytoplasm reddish blue, fine granules scattered, many in number in smear

Fills enlarged RBC, round or oval, compact cytoplasm, pale blue, Abundant brown granules

Smaller than RBC, very few in PBF, round compact, cytoplasm pale blue. Pigment and chromatin as in P. vivax

Of the size of RBC round, compact very few in PBF, cytoplasm pale blue, chromatin and pigment as in P. vivax

Host CELL

PARASITE General features

Delicate, small, 1.5 µm double chromatin and multiple rings common. Accole, wing and marginal forms

Lab Diagnosis of Malaria 209 Note: The lab diagnosis of filariasis is done by the peripheral blood smear examination. The methods mentioned in this chapter is applicable for the identification of microfilaria in peripheral blood.the sample collection is performed during night time. (Figs 53.10 and 53.11)

Figure 53.9: Parts of malaria parasite inside a RBC

Figure 53.10: Microfilaria in a blood smear

Figure 53.11: Structure of microfilaria

54 Parasitological Examination of Faeces ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

The examination of faeces for parasitological diagnosis is done to detect: Adult worms Segments of tapeworms Ova and cysts Larvae Trophozoites Cellular exudates such as WBCs, RBCs, macrophages and Charcot-Leyden (CL) crystals. Collection of Faecal Sample Ask the patient to pass the stool sample directly into a waxed cardboard or a plastic cup with a tight fitting lid. Collection of sample in a match box or on plant leaves is not a satisfactory method. About 20-40 grams of well-formed stool or 5-6 table spoonfuls of watery stool will suffice for a routine examination. Ingestion of some medicines prior to collection of faecal sample may interfere with the detection of parasites. These include tetracyclines, sulfonamides, antiprotozoal agents, laxatives, antacids,

Parasitological Examination of Faeces 211 castor oil, magnesium hydroxide, barium sulphate, bismuth kaolin compounds and hypertonic salts, etc. These should not be taken 1-2 weeks before the examination of stool sample. All specimens must be properly labelled with patient’s name, age, sex, and date of collection. The specimen must reach the laboratory within 30 minutes of passing of the stool, since amoebic trophozoites die and become unrecognizable after that. Note Do not keep the specimen at warm temperatures. Try to keep it in cool, shady places. Prevent the drying of the specimen. Prevent contamination with urine or dirt particles. Multiple stool examinations are required before the presence of parasitic infections is ruled out. Stool should not be collected from bed-pans containing disinfectants. Transportation of Samples If looking for trophozoites, stool specimen must be transported very rapidly to the laboratory to avoid disintegration of trophozoites. Stool samples should be examined within 30 minutes of collection of the specimen and not receipt of the specimen in the laboratory. Stool specimens should never be frozen and thawed or placed in an incubator because parasitic forms deteriorate very rapidly. For permanent fixation of the stool specimen, 10% formol-saline (prepared by adding 100 ml formaldehyde to 900 ml of 0.85% sodium chloride) is a well known preservative. Polyvinyl alcohol (PVA) is a widely used preservative because the performance of concentration procedures and preparation of permanent stained smears are both possible with this. Macroscopic Examination Various points to be noted are:

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Consistency: The consistency of the stool could be formed, soft, loose or watery. The cysts are found maximum in the formed stool while trophozoites are most abundant in watery stool. Presence of blood and mucus. Presence of round worms, thread worms or tapeworm proglottids. Colour and smell of the stool. Microscopic Examination (Temporary Wet Mounts) It is the simplest and easiest technique. A wet mount can be prepared directly from faecal material or from the concentrated specimens. The basic types of wet mounts that should be made from each sample include: a. Saline wet mount: It is used to detect worm eggs or larvae, protozoan trophozoites and cysts. In addition it can reveal the presence of RBCs and WBCs. b. Iodine wet mount: It is used to stain glycogen and nuclei of the cysts. Procedure Place a drop of saline on left half of the slide and one drop of iodine on the right half. With an applicator stick pickup a small portion of the specimen (equivalent to the size of a match head) and mix with saline drop. Similarly pickup similar amount and mix with a drop of iodine. Put the cover slip separately on both and examine under the microscope. Ova, cysts, trophozoites and adult worms can be identified as per their characteristic features (Figs 54.1 and 54.2). Iodine wet mount is examined for amoebic and flagellar cysts. Concentration Techniques If the number of parasites in the stool specimens is low, examination of a direct wet mount may not detect them, hence the stool should be concentrated. Eggs, cysts and larvae are recovered after concentration procedures whereas trophozoites get destroyed during the

Parasitological Examination of Faeces 213

Figure 54.1: Artifacts present in stool

Figure 54.2: Various structures seen in stool preparation

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procedure. This makes direct wet mount examination obligatory as the initial phase of microscopic examination. The concentration procedures can be grouped under 2 categories: a. Sedimentation procedures: In which the eggs and cysts settle down at the bottom. Examples. 1. Formol- ether sedimentation technique 2. Simple gravity sedimentation technique. b. Flotation procedures: In which the eggs and cysts float at the surface due to specific gravity gradient. Examples 1. Saturated salt solution technique 2. Zinc sulphate floation technique 3. Sugar flotation technique. 4. Saturated Magnesium sulphate solution technique. The basic disadvantage of sedimentation technique is that examination of the sediment is often difficult due to the presence of excessive faecal debris that may mask the presence of the parasites. The basic disadvantage of flotation technique is that not all eggs and cysts float in the flotation procedures. Two commonly used concentration techniques are formalin-ether and saturated salt solution technique. Formal ether sedimentation technique Procedure (Fig. 54.3) Transfer half teaspoonful of faeces in 10 ml of water in a glass container and mix thoroughly. Place 2 layers of gauze in a funnel and strain the contents into a 15 ml centrifuge tube. Centrifuge for 2 minutes at about 500 gm. Discard the supernatant and resuspend the sediment in 10 ml of physiological saline. Centrifuge at 500 gm and discard the supernatant. Resuspend the sediment in 7 ml of 10% formaldehyde (1 part of 40% formalin in 3 parts of saline). Add 3 ml of ether (or ethyl acetate). Close the tube with a stopper and shake vigorously to mix. Remove the stopper and centrifuge at 500 gm for 2 minutes.

Parasitological Examination of Faeces 215

Figure 54.3: Formol-ether sedimentation technique

Rest the tube in a stand. Four layers now become visible the top layer consists of ether, second is a plug of debris, and third is a clear layer of formalin and the fourth is the sediment. Detach the plug of debris from the side of the tube with the aid of a glass rod and pour off the liquid leaving a small amount of formalin for suspension of the sediment. With a pipette, remove the sediment and mix it with a drop of iodine. Examine under the microscope. Advantages Faecal odour is removed. The sensitivity of detecting the ova or cysts increases by 8-10 folds. The examination is easier than examining a direct wet smear. The size and shape of the parasitic structures is maintained. It is inexpensive, easy to perform and can be done at any level of health infrastructure. Disadvantages Faecal debris may mask the parasitic structure. Trophozoite forms are not detected in this method.

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Saturated Salt Flotation Technique Place about one millilitre of faeces in a container which is flat bottomed and has a diameter of less than 1½ inches and capacity of about 15-20 ml (Fig. 54.4).

Figure 54.4: Salt floatation technique

Add a few drops of saturated salt solution (specific gravity 1.200) and stir it to make a fine emulsion. Add more salt solution so that the container is nearly full, stirring the solution throughout. Remove any coarse matter which floats up. Place the container on a level surface. Do the final filling by a dropper until a convex meniscus is formed. A glass slide 3" × 2" is carefully laid on the top of the container so that the centre is in contact with the fluid. Preparation is allowed to stand for 20 minutes after which the glass slide is quickly lifted, turned over smoothly as to avoid spilling of the fluid and examined under the microscope after putting a cover slip. Disadvantages of Flotation Techniques Unfertilized eggs of Ascaris lumbricoides, eggs of Taenia solium and Taenia saginata, all Trematodal eggs and larvae of Strongyloides do not float in the salt solution.

Parasitological Examination of Faeces 217 Due to high specific gravity of the solution, protozoan cysts and thin walled nematode eggs will collapse and become distorted in appearance, if left for more than 20 minutes. Reporting of Results (Figs 54.5 to 54.9) The report should include positive/negative comments on the following: • Adult worms/segments of worms/larvae. • Cellular exudate such as RBCs, WBCs, Macrophages and CL crystals. • Trophozoites (only if the specimen was fresh otherwise comment that specimen was not fit to comment on this). • Ova and cysts. Any advice for further examination.

Figure 54.5: Trophozoites and cysts in stool preparation

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Figure 54.6: Hookworm larvae and egg

Figure 54.7: Whipworm egg and adult worms

Figure 54.8: Pinworm egg and adult worms

Parasitological Examination of Faeces 219

Figure 54.9: Roundworm eggs and adult worms

55 Medical Entomology ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Insects and arachnids belong to the Phylum Arthropoda. Insects and arachnids are associated with many parasitic infections and other microbial diseases. Knowledge of their characteristics, breeding and feeding habits are useful to the prevention and control of the diseases they produces. The study of insects is known as entomology. The infections transmitted by these agents are called as arthropod borne or vector borne infections. The insects may act either as Biological vector, carry the microorganism in their body and pathogen undergoes some development in the vector or as Mechanical vector just carry the pathogen in their body and pathogen doesnot under go any development in the vector. Biological vector example: Female Anopheles mosquito for malarial parasite. Mechanical vector example: House flies for bacillary dysentery pathogen (Shigella). The body of an insect is divided in to head, thorax and abdomen. The head bears mouth parts and a single pair of antennae. Thorax bare three pairs of legs, many insects are equipped with two pairs of wings.

Medical Entomology 221 Insects

Arachnids -

Mosquitoes Fleas Bugs Lice Ticks Mites

Mosquitoes Mosquitoes are the most wide spread medically important insects. Female mosquitoes need blood for egg production so, they bite and suck blood from human or animals. Female mosquitoes are thus responsible for transmission of disease. Male mosquitoes do not bite and therefore they dot not transmit disease. They live on plant juice. The life span of an adult mosquito is generally 3-4 weeks (Fig. 55.1).

Figure 55.1: Developmental stages of anopheline and Culex mosquitoes

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Classification of Mosquitoes The mosquitoes of medical importance are: – Anopheles – Aedes – Culex – Mansonia Anopheles Mosquitoes (Fig. 55.2) 1. Diseases transmitted: – Malaria – Bancroftian and Burgian filariasis – Viral encephalitis (arbovirus) 2. Feeding habits: – Night feeders – Anopheline mosquitoes rest with the body sloping forwards 3. Breeding sites: – Pools – Swamps – Seepages – Rice fields – Tree – holes – Ditches

Figure 55.2: Anopheles mosquitoes

Medical Entomology 223 Aedes Mosquitoes (Fig. 55.3) 1. Diseases transmitted: • Bancroftian filariasis • Yellow fever • Dengue • Chickun gunya • Rift valley fever 2. Feeding habits: • Feeding during the day or night indoor and out door 3. Breeding sites • Stagnant water in tree holes • Coconut shells • Old tins • Pots

Figure 55.3: Aedes mosquitoes

Culex Mosquitoes (Fig. 55.4) 1. Diseases transmitted • Bancroftian filariasis • Japanese encephalitis • West Nile fever • Western Equine encephalitis 2. Feeding habits • Mostly indoor night feeder • Attracted to animals

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Figure 55.4: Culex mosquitoes

3. Breeding sites • Latrines • Waste water Mansonia Mosquitoes (Fig. 55.5) 1. Disease transmitted: Brugian filariasis Bancroftian filariasis 2. Feeding habits Night feeders of indoors and out door 3. Breeding sites Paddy fields, swamps, collections of water containing water plants or swamp grass.

Figure 55.5: Mansonia mosquitoes

Medical Entomology 225 Sandy Fly (Phlebotomus spp) (Fig. 55.6) Diseases Transmitted • Visceral Leishmaniasis (Kala-azar) • Cutaneous Leishmaniasis • Sandfly fever • Oroya fever (Carrion’s disease) Morphology • Small hairy, yellow or grey colored insect • 3-5 mm in length • Long antennae • Upward held wings. • Phlebotomus species are found in warm and hot climates like desert. They fly near to the ground.

Figure 55.6: Sand fly

Feeding Habits The female sucks blood from humans’ rodents, dogs and other animals, male feeds on plant juices. Breeding The eggs are load in cracks and holes in the moist ground. There are four larval stages before pupae are formed. The adult sand fly lives for few weeks.

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TseTse Flies (Glossina palpalis) (Fig. 55.7) Disease transmitted African trypanosomiasis (sleeping sickness) Morphology • Tsetse flies are large yellow – brown or brown – black • 6-15 mm in length • They have long a proboscis and short antennae. • Axe shaped venation in the wings.

Figure 55.7: Tsetse fly

Feeding Habits Both male and female suck blood. They feed on human beings and other mammals. Breeding Sites and Life Cycle • G.palpalis lives in vegetation bordering rivers, lakes and other water places. • A tsetse fly does not lay eggs but produces a single fully developed larva. • The life span of a tsetse fly is 2-3 months. HOUSE FLY (MUSCA DOMESTICA) (FIG. 55.8) Diseases Transmitted (Mechanical vector) • Typhoid • Bacillary dysentery

Medical Entomology 227

Figure 55.8: House fly

• Cholera • Plague • Brucellosis • Tetanus • Amoebic dysentery • Streptococcal infection • Polio myelitis • Hepatitis A Morphology • 8 mm length • Greyish black with four dark stripes on its thorax • The antennae are short • Proboscis is retractable Feeding Habit • Feed during the daytime • Non-biting Breeding and Life Cycle • Eggs are laid in large numbers in manure and other waste matter. • In warm conditions, the larvae (Maggots) hatch • Adult live for 1 month. RAT FLEA (XENOPSYLLA) (FIG. 55.9) Diseases Transmitted • Bubonic plague • Flea-borne murine typhus (Endemic typhus)

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Figure 55.9: Rat flea

Species • X. Cheopis • X. asiatica • X. brasilliensis Feeding Habit • Feed on blood of mammals and birds • Infestrats. • When rats are killed by the disease (Plague) the fleas leave the rats and can become parasitic on human beings. • Female ingests more blood than the male Breeding The female fleas lays eggs in batches in or near the rat. There are three larval stages and a pupal stage. • Adult flea can live upto 5 years. • Unfed flea can live only upto 1 year. LOUSE (PEDICULUS HUMANUS) (FIGS 55.10 AND 55.11) Species • P. humanus var capitis (head louse) • P humanus var corporis (body louse) Diseases Transmitted • Relapsing fever (louse borne) • Epidemic typhus (louse borne) • Trench fever. • Louse bites also cause irritation and scratching which lead to secondary infection.

Medical Entomology 229 Morphology • 2-3 mm in length. • Head is small and bears a pair of short antennae, small eyes and mouth parts adapted for sucking. • Thorax without wings. The legs are well developed for gripping.

Figure 55.10: Head louse

Figure 55.11: Crab louse

Feeding and Breeding • Lice are ectoparasites of humans of animals. • Life cycle is a incomplete metamorphosis. • Eggs are laid on hair or clothing and fixed firmly to the host by a glue like substance secreted around the eggs by female. Both sexes feed on blood. ITCH MITE (SARCOPTES) (FIG. 55.12)

Species: Sarcoptes scabiei Disease produced: Scabies The female mite burrows into the skin to lay its eggs. This usually occurs where the skin is thin and in children. Penetration of the mite causes severe irritation and a rash. Scratching may lead to secondary infection.

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Figure 55.12: Itch mite

Morphology Very small 0.3 mm in diameter Microscopical examination of the skin scrapping of the suspected area may show the parasites and it’s eggs. TICKS Ticks of medically importance belong to two groups, hard ticks and softticks. Hard Ticks They belong to Ixodidae family (ixodid ticks) Important genera • Dermacentor • Amblyomma Disease Transmitted • Tick-borne typhus (Amblyomma) • Tick borne encephalits (Dermacentor) • Q fever and Tularemia are also thought to transmitted by hard ticks. Morphology • Mouth parts project in front • 4-5 mm in length

Medical Entomology 231 • Dark or brightly coloured with metallic markings • Hairs usually absent. Feeding and Breeding • Parasitise humans and animals • Both sex feed as blood. The eggs are laid in a single batch after which the female dies. Each stage of development (larvae- lymph) may be spent on a different host. Soft Ticks (Argasid ticks) They belong to Argasidae family important species are: • Ornithodoros moubata. • Haema physalis splinlgera Disease Transmitted • Borrelia Relopsing fever (Onithodoros) • Kyasanur Forest disease (Haemaphysalis) Morphlogy • 5 mm length • Oval in shape • Short legs and from above no mouth parts visible • Hairs usually absent Feeding and Breeding • Ectoparasite of man and monkeys. • Soft ticks can survive for long periods without host. • Both sexes suck blood from man. • The eggs are laid in batches in cracks and crevices of walls. The life cycle takes 6-12 months. • Soft ticks survive best in dry climates. CYCLOPS (WATER FLEAS) (FIG. 55.13) Water Fleas belong to the class Crustacea, which also include crabs, prawns and other sea and fresh water creatures. Disease Transmitted • Guinea worm infection • Diphyllobothrium latum infection. • Cyclops is the intermediate host of helminthic parasites Diphyllo bothrium latum and Dracunculus medinensis (Guinea worm).

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Figure 55.13: Cyclops

Morphology • A Tiny arthropod, not more than 1mm in length and just visible to the naked eye. • Pear–shaped semitransparent body • Forked tail • 2 pairs of antennae • 5 pairs of legs and a small pigmented eye. Feeding and Breeding • It swims in fresh water with jerky movements. • The average life span of a cyclop is about 3 months. • Each cyclop can ingest 15-29 embryos of Guinea worm. Hyperinfection of cyclops can reduce their growth or even kill it. • The embryos feed on gonards of cyclops. • It takes 2 weeks for the development of Guinea worm larvae in cyclops. Control of Guinea worm infection by ingestion of cyclops Filtering drinking water Boiling all drinking water Avoid step-well contamination. Use organophosphorus chemicals or chlorine to destroy cyclops. Biological control of Cyclops can be done by Barbel fish and Gambusia fish.

56 Bacteriological Examination of Water ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Water is the elixir of life. Water of adequate quantity and quality, is essential for healthy life. Many diseases are associated with contaminated water. Contamination of water results in various parasitic infestations, fungal skin diseases, eye infections and diarrheal diseases like cholera, dysentery and enteric fever. Microbiological examination offers the most sensitive test for the detection of recent and potentially dangerous faecal pollution, thereby providing a hygienic assessment of water quality with high sensitivity and specificity. It is ideal to look for individual specific pathogen but it is not practical since they are few in numbers than the nonpathogenic organisms and methods to detect them are costly in time and money. Therefore indicators of human/animal pollution, e.g. coliforms are used. Faecal Streptococci are regularly present in the faeces in varying numbers but their number is fewer than Esch. coli and they probably die and disappear at the same rate. The presence of faecal Streptococci along with coliforms in absence of Esch. coli is also confirmatory of faecal pollution (Fig. 56.1). Water Sampling Sources of water to be sampled:

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Figure 56.1: Dilution of water for coli-form count

Water sources can be divided into three basic types for the purpose of sampling. a. Water from a tap or fixed hand pump b. Water from a reservoir (river, lake, and tank) c . Water from a dug well

Bacteriological Examination of Water 235 Sampling from a Tap or Pump Outlet a. Remove any attachments from tap that may cause splashing. b. Wipe off the dirt from outside the tap. c . Turn on the tap at maximum flow rate and let the water flow for 12 minutes. d. Sterilize it for a minute with flame using gas burner, lighter or ignited cotton wool soaked in spirit. e . Open the tap and allow water to flow at medium rate for 1-2 minutes. . f Open the container for collecting the sample and fill the water by holding the bottle under the water jet. Leave a small airspace to facilitate shaking at the time of inoculation prior to analysis. g. Stopper the cap and label the container. Sampling from Reservoir a. Open the bottle under sterilized conditions. b. Fill it by holding it by the lower part, submerging it to a depth of about 20 cm, with the mouth facing slightly upwards. If there is a current, the bottle should face the current. c . Stopper the bottle and label it. Sampling from a Dug Well a. Attach a stone of suitable size to the sampling bottle with a piece ofstring. b. Tie a 20 meter length of clean string on the bottle and to a stick. c . Open the bottle as described above and lower into the well. d. Immerse the bottle completely in water without touching the sides of the well and lower it down to the bottom of the well. e . Pull it out when the bottle is filled. . f Discard a little water to provide airspace. g. Stopper and label the bottle. The sample is examined at the earliest preferably within one hour or should be quickly transported to the laboratory keeping in cool container away from sunlight. It should be positively examined within 6 hours of collection.

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While sampling chlorinated water 0.5 ml of sodium thiosulphate solution (18 gm/L) should be added to sampling bottles to neutralize the residual chlorine present in water. Methods of Analysis Two methods have been developed for the detection of indicator bacteria in water: multiple tube method and membrane filter method. Multiple-tube method. In the multiple-tube method different amounts of water to be tested are added to tubes containing a suitable culture medium. The bacteria present in the water reproduce and produce acid with or without gas. From the number of tubes inoculated and the number with a positive reaction, the most probable number (MPN) of bacteria present in the original water sample can be determined statistically. The multiple-tube method is applicable to all kinds of water: it can be used with clear, colored, or turbid water containing sewage or sewage sludge, or mud and soil particles, provided that the bacteria are homogeneously distributed in the prepared test samples. Procedure Set up following volumes of different strengths of MacConkey broth in tubes/bottles each having an inverted Durham tube to detect the presence of gas and add specified volume of water as mentioned: • One 50 ml of water to 50 ml of double strength medium. • 5, 10 ml quantities each to 10 ml double strength medium. • 5, 1 ml quantities each to 5 ml single strength medium. • Incubate the tubes/bottles at 37oC for 18-24 hours. Observe change in color and appearance of gas in Durham tubes inbottles. The media receiving one or more of the indicator bacteria show growth and a color change which is absent in those receiving an inoculum of water without indicator bacteria. Presence of both acid and gas indicates positive reaction whereas absence of either or both these features denotes a negative reaction. The presumptive positives are read and remaining negative bottles are re incubated for another 24 hours. Any further positives

Bacteriological Examination of Water 237 are added to the previous figures. The probable number of coliforms is read from the probability tables of McCrady. From the number and distribution of positive and negative reactions, count of the most probable number (MPN) of indicator organisms in the sample may be estimated by reference to statistical tables. The test gives presumptive coliform count as the reaction observed may occasionally be due to the presence of some organisms other than coliforms. Most probable number of coliforms by McCrady’s table

No. of tubes giving positive reactions 1×50 0 0 0 0 0 0 0 0 0 0 0 0 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1

— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

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— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

ml————

5×10

— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

0 0 0 1 1 1 2 2 2 3 3 4 0 0 0 0 1 1 1 1 2 2 2 2 3 3 3 3 3 4 4 4 4

— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

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— — — — — — — — — — — — — — —

— — — — — — — — — — — — — — — — —

— — — — — — — — — — — — — — — — —

ml———— 5×1 — — — — — — — — — — — — — — —

— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — ——————— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

0 1 2 0 1 2 0 1 2 0 1 0 0 1 2 3 0 1 2 3 0 1 2 3 0 1 2 3 4 0 1 2 3

— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

ml———— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

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— — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — — —

MPN/100 ml— < 1 2 1 2 3 2 3 4 3 5 5 1 3 4 6 3 5 7 9 5 7 1 1 8 1 1 1 2 1 1 2 2

1 — — — — — — — — — — — — — — — — — — — — — 0 2 1 4 8 1 3 7 2 8

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Contd... 1 1 1 1 1 1 1 1 —

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4 4 5 5 5 5 5 5 —

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4 5 0 1 2 3 4 5 —

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35——————— 43——————— 24——————— 35——————— 54——————— 92——————— 161——————— >180—————— — — — — — — —

Eijkman Test (differential coliform test or confirmed E.coli count) Eijkman test is usually done to confirm that the coliform bacilli detected in the presumptive test are E. coli., as some spore – bearing bacteria give false-positive reactions in the presumptive coliform test. After the presumptive test, subcultures are made from all tubes showing acid and gas to fresh tubes of single strength MacConkey medium which is brought to 37oC. These tubes are incubated at 44oC in thermostatically controlled water baths and examined after 24 hours. Those tubes showing acid and gas are that containing E. coli, the number is read from the McCrady table. E.coli can be confirmed by plating on solid media and testing for indole production and citrate utilisation. Standards The classification of drinking water according to bacteriological tests is given below: Class

Grade

I Excellent

Presumptive count (per 100 ml)

E.coli count (per 100 ml)

0

0

II

Satisfactory

1-3

0

III

Suspicious

4-10

0

IV

Unsatisfactory

>10

0,1 or more

Reporting Mention the presumptive coliform count and Escherichia coli count per 100 ml of water. Report: Fit/unfit for human consumption.

57 Microbiology of Milk ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

INTRODUCTION Milkborne infections can be prevented by proper sterilisation of milk. Milk is commonly sterilised by pasteurisation or boiling. Infections of animals can be transmitted to human beings through contaminated milk. Examples are Brucellosis, Bovine tuberculosis, Salmonellosis and Q-fever. Very merely Anthrax and Leptospirosis can also be transmitted through milk. The routine bacteriological examination of milk especially in diary farms can prevent the above mentioned diseases. Common test performed for bacteriological examination of milk include. 1. Viable count: Raw milk always contains bacteria, varying in number from 500 to several million per ml of milk. This can be detected by plate counts. 2. Test for coliform bacteria: Contamination of milk may show the presence of coliform bacteria. This can be detected by inoculating milk in to Mac Conkey’s fluid medium. 3. Mehtylene blue reduction test: This is a simple test for the viable count. It depends on the reduction of methylene blue by bacteria in milk when incubated at 37oC in complete darkness. The rate of reduction is related to the degree of bacterial contamination. The change of blue color to colourless indicates the reduction.

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4. Phosphatase test: This is a test for pasteurization of milk. The enzyme phosphatase normally present in milk is inactivated if pasteurization has been carried out properly. Residual phosphatase activity shows that pasteurization has not been adequate. 5. Turbidity test: If milk is boiled to the prescribed temperature and time, all heat coagulable proteins are precipitated. If ammonium sulphate is then added to the milk, filtered and boiled for 5 minutes, no turbidity results. This test can be used differentiate pasteurised and sterilised milk.

58 Lab Diagnosis of Tuberculosis ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Tuberculosis is caused by Mycobacterium tuberculosis which is an acid fast bacillus (AFB). The highest priority for tuberculosis (TB) control is the identification and cure of infectious cases, i.e. patients with sputum smear positive pulmonary TB. The highest priority in the diagnosis of TB is thus given to sputum microscopy. Morphology of M. Tuberculosis Acid fast bacilli are approximately 1-10 m long, slender, rod-shaped bacilli which may be curved or bent. These may be granular, isolated, in pairs or in groups. Stained bacilli may present a beaded appearance. Microscopy of Sputum Diagnosis of pulmonary TB by sputum microscopy is simple, easy, inexpensive, rapid, technically not very demanding and more reliable than X-rays. The purpose of the sputum microscopy is helpful for (a) diagnosis of the patients with infectious tuberculosis (b) monitoring the progress. For diagnosis, 3 sputum examinations are performed (spot, morning, spot) and for follow up 2 sputum examinations (morning, spot) are performed.

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Collection of Sputum Sample • Select a good wide-mouthed sputum container, which is disposable, made of clear thin plastic, unbreakable and leak proof material. • Give the patient a sputum container with the laboratory serial No. written on it. Show the patient how to open and close the container and explain the importance of not rubbing off the number written on the side of the container. • Instruct the patient to inhale deeply 2-3 times, cough up deeply from the chest and spit in the sputum container by bringing it closer to mouth. • Make sure the sputum sample is of good quality. A good sputum sample is thick, purulent and sufficient in amount (2-3 ml). Give the patient another container with laboratory serial number written on it for an early morning specimen. Explain to the patient to rinse his/her mouth with plain water before bringing up the sputum. Storage and Transportation of Specimens If the specimen is collected in the field and cannot be immediately processed, it should be transported to the laboratory within 3-4 days of collection. The specimen should be collected in the containers meant for the purpose, lid tightly secured, properly labelled and kept away from the sun and heat. These can be placed in a special box which can withstand leakage of contents, shocks and other conditions incident to ordinary handling practices. These boxes should be kept in the cooler conditions and then transported to the laboratory. Preparation of Smear and Ziehl Neelsen Staining (AFB staining) • Select new unscratched slide and label the slide with Laboratory Serial number. • Make a smear from yellow purulent portion of the sputum using a thin stick. A good smear is spread evenly, 2 cm × 3 cm in size and is neither too thick nor too thin. The optimum thickness of the smear can be assessed by placing the smear on a printed matter and the print should be readable through the smear.

Lab Diagnosis of Tuberculosis 243 • Let the smear air dry for 15-30 minutes. • Fix the smear by passing the slide over the flame 3-5 times for 3-4 seconds each time. • Stain the smear by Ziehl Neelsen method. Grading of Microscopy Smears Record the results in laboratory form and laboratory register appropriately as per table given below: Examination

Result

Grading

No. of fields to be examined

More than 10 AFB per oil immersion fields

Positive

3+

20

1-10 AFB per oil immersion fields

Positive

2+

50

10-99 AFB per 100 oil immersion fields

Positive

1+

100

1-9 AFB per 100 oil immersion fields

Scanty

Record exact number seen

100

No AFB per 100 oil immersion fields

Negative

0

100

How to Prevent False Positive Sputum Results? • • • • • • • • • •

Only use new, unscratched slides. Always use filtered carbol fuchsin. Do not allow the carbol fuchsin to dry during staining. Do not allow the carbol fuchsin to boil during staining. Decolorize adequately with sulphuric acid. Make sure there are no food particles or fibres in the sputum sample. Never allow the oil immersion applicator to touch a slide. Never allow the oil immersion lens to touch a slide. Label sputum containers, slides, and laboratory forms accurately. Record and report results accurately.

How to Prevent False Negative Sputum Results? • Make sure the sample contains sputum and not just saliva.

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• Make sure there is enough sputum (at least 2 ml). • Select thick, purulent portions to make the smear. • Prepare smears correctly – not too thick, too thin or with too little material. • Fix for the correct length of time, not too short or too long. • Stain with carbol fuchsin for 5-7 minutes. • Do not decolorize with sulphuric acid too intensively. • Examine every smear for at least five minutes observing at least 100 fields before recording as negative. • Label the sputum containers, slides and laboratory forms carefully. • Record and report results accurately. Culture for M. tuberculosis

Indications Each and every case of suspected tuberculosis need not be cultured. The judicious use of culture limits its use to the followings: • Diagnosis of smear negative pulmonary TB cases with strong clinical and radiological suspicion. • Diagnosis of extrapulmonary TB. • Follow up of a case to investigate for a drug resistant isolate. • Diagnosis of childhood tuberculosis.

Procedure In some of the intermediate laboratories, the culture for M. tuberculosis may be feasible – wherever indicated and possible. The procedure recommended is given below: • In a small glass, sealable bottle, mix equal volumes of sputum and 4% sodium hydroxide. • Shake well and incubate at room temperature (25-30oC) for 15-20 minutes with regular shaking every 5 minutes. • Centrifuge at high speed (> 13,000 gm) for 8-10 m. • Discard the supernatant. • Neutralize the sediment by adding drop by drop, 2 mol/litre HCl containing 20 ml of phenol red solution per litre until the mixture remains pink.

Lab Diagnosis of Tuberculosis 245 • Inoculate the neutralized deposit on to atleast 3 tubes of Lowenstein-Jensen (L-J) medium-one with glycerol and another with sodium pyruvate (Fig. 58.1). • Incubate the L-J tubes for 2-3 days at 35-37oC in a horizontal position with the caps loosened half a turn. Thereafter incubate at 37oC for 6 weeks and inspect for growth at weekly intervals. Initial incubation of the culture tubes in the presence of 5-10 percent CO2 improves the growth of M.tuberculosis. • Note the growth of bacteria on the surface during these weekly inspections, and if present stain by Ziehl-Neelsen method. • If the isolate has the typical colonial appearance and the Z-N stained smear from a colony is also typical report the growth as Mycobacterium spp. • Send the isolate to a reference laboratory for further characterisation and susceptibility testing. The growth of typical human strains is "rough, tough and buff" and seen after 2-3 weeks of incubation but seldom earlier.

Figure 58.1: L-J medium with M.tuberculosis colonies

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Biosafety • Treat all sputum samples as potentially infectious and use leak proof containers for collection and transportation. • Use bacteriological safety hood while carrying out all procedures involving sputum. • Dispose of the sputum cups by incineration, autoclaving or treating with 5% phenol or 2% freshly prepared hypochlorite solution, whichever is feasible. • Take special precautions while transporting the cultures to the reference laboratories with emphasis on containers which don’t get broken in the event of an accident. • Wash hands with soap and water frequently especially after touching the sputum. Reporting of Results Sputum microscopy • If all three specimens or two specimens show presence of AFBs: Report positive. • If one of three specimens show AFB, correlate the findings clinically and radiologically. • If none of the three specimens shows AFBs: Report negative. Positive culture • Report positive when typical colonies are isolated and Ziehl Neelsen staining shows acid fast bacilli. Negative culture If no growth is obtained after 6-8 weeks of incubation report negative.

59 Urinary Tract Infection ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

Urinary Tract Infections (UTI) could be of the lower urinary tract encompassing the bladder and urethera or of the upper urinary tract infecting the ureters and kidneys. Because of shorter urethera, bacteria can reach the bladder more easily in females. All areas of the urinary tract above the urethera in healthy humans are sterile, hence urine is normally sterile. UTI is among the most common reasons patients seek medical care. It is estimated that approximately 10% of humans will have UTI at sometime during their lives. Causative Organisms

Escherchia. coli is by far the commonest cause of uncomplicated community acquired urinary tract infections. Other members of Enterobacteriacae can also infect. The causative agents are listed below. • E.coli • Proteus mirabilis • Staph. aureus • Staph. saprophyticus • Group B streptococci • Enterococci

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Complicated UTI which occurs in cathetrised patients or with obstruction is caused by: • E.coli • Klebsiellaspp. • Enterobacter • Proteus spp. • Pseudomonas aeruginosa • Staph. aureus Specimen Collection Prevention of contamination by normal vaginal, perineal and anterior uretheral flora is very vital. It is the responsibility of the laboratory to provide the patient with sterile, wide-mouthed glass or plastic, jars, beakers or other suitable receptacles which should have tight-fitting lids. Though urine collected by suprapubic aspiration is the gold standard, it is not a practical method. Alternatively, mid-stream urine or a clean catch urine is collected. Whenever possible, urine specimen should be collected in the morning, before the patient has voided urine. The collection method is described as follows: In Men Instruct the patient to wash hands. Ask the patient to pull back the foreskin and pass a small amount of urine holding back the fold of skin, instruct the patients to pass the remaining urine in a sterile container–this is mid-stream urine (MSU). Place the lid, secure tightly and rapidly transport to the laboratory. If the patient is bed-ridden, the nursing staff can help the patient in the above process. The sample is best obtained, if the procedure is properly explained to the patient.

Urinary Tract Infection 249 In Women Instruct the woman to wash hands with soap and water before collection of specimen. Patient should undress in a suitable room, spread the labia and cleanse the vulva and labia thoroughly using warm soapy water. Rinse with warm water and dry. Ask the patient to pass urine, discarding the first part of the stream and collecting MSU in a sterile container. Transport the sample to the laboratory at the earliest after properly securing the lid. Bed- ridden patients can be assisted by the nursing staff. In Infant and Young Children Ask the child to drink water or any other liquid. Clean the external genitalia and let the child be seated in the lap of the mother/nurse/attendant. Encourage the child to urinate and collect the same in sterile container. Cover the container tightly and rapidly transport to the laboratory for processing. CoIlection of urine from catheters or bag should be avoided as this does not reflect the accurate picture. A reasonable alternative to MSU is the clean catch urine. After periuretheral cleaning the whole urine is collected into a sterile container and then an aliquot is sent for examination. Specimen Transport Pending immediate processing of the sample, urine should be refrigerated. Bacterial counts at 4oC remain constant for 24 hours. Beyond 48 hours, even the refrigerated samples are not suitable. Salient features of specimen collection Suprapublic aspiration of urine sample which is the best is not always possible. Mid Stream Urine and Clean Catch Urine are satisfactory alternatives.

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Process the sample at the earliest preferably within 4 hours. If immediate processing is not possible, refrigerate the sample at 4oC, upto a maximum of 48 hours. Catheter urine and bag urine are not ideal samples. Screening Procedures Upto 80% of the urine specimen received in laboratory for culture may contain no etiological agent of infection or may contain only contaminants. There are various procedures tried to screen out such samples so that time, reagents and money of the laboratory is saved. Of these, a simple Gram stained smear of the urine has been found to be least expensive and probably the most sensitive and reliable screening method. Gram stained smear of urine Place a drop of well mixed urine sample on a clean, grease-free slide. Air dry and heat fix. Perform Gram staining. Examine under oil immersion (100x). Presence of atleast one organism per oil immersion field (examine at least 20 field) correlates with significant bacteriuria (> 1 × 105/ml). Culture Urine is cultured in the following situations Sample screened by Gram stain and found to have significant bacteriuria. Follow up of patients on treatment. Urinary tract obstruction. Follow up after removal of indwelling catheter. Bacteremia of unknown origin. Once it has been decided that urine sample is to be cultured, a measured amount is inoculated to each of the appropriate media (Fig. 59.1). Usually a calibrated loop designed to deliver a known volume (say 0.01 ml per loopful) of urine is used. The method followed is as follows:

Urinary Tract Infection 251

Figure 59.1: Inoculation of urine sample

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Mix the urine sample thoroughly and remove the top of the container. Flame a calibrated wire inoculating loop and allow it to cool without touching any surface. Insert this sterile loop in the urine sample vertically and allow urine to adhere to the loop. Spread the loopful of urine over MacConkey agar plate using standard method. In the similar way, collect a second loopful specimen and inoculate a blood agar plate. Incubate the plates aerobically at 35-37oC for at least 24 hours. Count the colonies and count the CFU (colony forming unit) by multiplying with 100 (since 0.01 ml loop was used). Reporting of Results Report on number of WBC, RBC/ml of urine. Comment on the presence of epithelial cells, yeasts or parasites (e.g. Trichomonas vaginalis). If culture not indicated report "Culture not done because counts below signficant levels". Report on the type of organism isolated alongwith counts – significant or otherwise. Report absence of growth. Report susceptibility results as clinically indicated.

60 Spotters ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

PLATINUM WIRE LOOP (FIG. 19.4) 1. Used to pick up colonies grown on culture media and to culture the clinical specimens. 2. Sterilised by heating in a Bunsen flame till it becomes red hot. CANDLE JAR (FIG. 20.2) 1. The candle jar is employed to provide 5-10% CO2 atomosphere for the cultivation of carboxyphilic bacteria like: Pneumococci, Gonococci, and Meningococci, etc. 2. The inoculated culture plates are placed inside the jar and a candle is lighted inside. Then the lid is put on and made air-tight with Vaseline. The candle uses up all the oxygen present inside the jar and there will be accumulation of carbon dioxide. The whole jar is then incubated at 37oC and observed for growth. TESTING OF AUTOCLAVES 1. Spore indicator: A themophilic organism like Bacillus stearothermophilus with an optimum growth temperature of 5560oC is used whose spores require an exposure of 12 minutes at 121oC to be killed. Paper strips impregnated with spores are

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placed in paper envelopes and are inserted in different parts of a load : after sterilisation, the strips are inoculated into a suitable recovering medium and incubated for sterility test at 55oC/ 5 days. 2. These tubes contain an indicator liquid, which changes its colour from red to green, if the correct temperature time ratio, has been employed. If not they turn to reddish-brown colour. 3. Autoclave tapes 4. Thermocouples SEITZ FILTER 1. This is an asbestos disc filter and is most satisfactory for general purposes. 2. Seitz filter is attached to a vacuum pump through a silicone rubber bung. 3. Used to sterilise sera, sugar and antibiotic solutions. CENTRIFUGE (FIG. 22.2) 1. An electrically devised apparatus for the separation of two substances of different density by centrigugal force. Uses a. For the separation of particulate microorganisms from suspending fluid b. To obtain cell-free serum from clotted blood THROAT SWAB (FIG. 22.3) 1. This is used to swab the throat, wounds ,etc 2. It is sterilised by autoclaving or in hot-air oven. GLUCOSE BROTH 1. This is an enriched, liquid medium. 2. Widely used in blood culture. 3. It consists of nutrient broth with 0.5% sterile glucose.

Spotters 255 GELATIN 1. It is a protein derived from the collagen of skin and bone, etc. 2. Gelatin is used as a gelling agent. 3. It is also used to determine the proteolytic nature of bacteria and theirabilitytoliquifyit. 4. Its melting point is 22oC. NUTRIENT AGAR 1. It is a simple solid medium used for ‘the cultivation of nonfastidious organisms. 2. It consists of peptone, meat extract, sodium chloride and distilled water. 3. Also used to study the pigmentation, produced by bacteria. 4. Used in antibiotic sensitivity tests. BLOOD AGAR 1. It is an enriched medium, used for the isolation of fastidious organisms. 2. It consists of nutrient agar with 5-10% blood. AGAR AGAR 1. It is a solidifying substance, most widely used in the preparation of media. 2. Agar does not have any nutritional value. 3. It is derived from sea weeds. 4. The melting point of agar is 98oC and it solidifies at 42oC. SABOURAUD’S GLUCOSE AGAR 1. This is a solid medium used for the cultivation of different fungi. 2. It consists of peptone, glucose, agar and water. 3. The pH of the medium is 5.4.

STAPHYLOCOCCI (FIG. 60.1) 1. Gram Positive cocci.

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Figure 60.1: Gram’s smear of Staphylococci

2. Arranged mainly in groups, pairs and short chains are also found. 3. Cluster formation is due to cell division occuring in three planes, with daughter cells tending to remain in close proximity. 4. Non-sporing and non-capsulated. HAEMOLYSIS ON BLOOD AGAR 1. Haemolytic bacteria produce a definite zone of clearance around their colonies by lysing the red blood cells present in blood sgar. 2. Haemolysis is due to the production of an exotoxin called Haemolysin by the bacteria 3. Example of haemolytic bacteria; 1. Staph. aureus (Fig. 60.1). 2. Strep.pyogenes (Fig. 60.2).

Figure 60.2: Gram’s smear of Streptococci

Spotters 257 NEISSERIA GONORRHOEAE 1. Gram negative, bean-shaped diplococci with the adjacent sides concave. 2. Intracellular in positive being found within the pus cells. GRAM POSITIVE BACILLI 1. Violet coloured, rod shaped bacteria. 2. Found discretely and few in chains. Ex: Bacillus subtilis. THIOGLYCOLLATE MEDIUM 1. Used for the cultivation of anaerobes. 2. Contains mainly sodium thioglycollate, glucose and methylene blue. 3. Glucose acts as a reducing agent: thioglycollate maintains the anaerobic condition and methylene blue acts as an indicator. ROBERTSON’S COOKED MEAT MEDIUM 1. 2. 3. 4.

It is liquid anaerobic medium. Used for the cultivation of anaerobic organisms. It consists mainly of meat particles. The unsaturated fatty acids present in the medium absorb the oxygen within the bottle, the reaction being catalysed by haematin present in the meat particles, thus creating complete anaerobiosis. 5. Saccharolytic organisms turn the meat particles pink and proteolytic organisms turn the meat particles black. McINTOSH AND FILDE’S JAR 1. It is an anaerobic jar widely used for the cultivation of anaerobic microorganisms. 2. It consists of a stout metallic jar with a lid which carries gas inlet and outlet, two electrical terminals and a porcelain spool wrapped by palladium shed asbestos on the underside. 3. The sidearm carries a small tube containing reduced methylene blue indicator for verifying the anaerobic condition in the jars.

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MAcCONKEY’S AGAR WITH LACTOSE AND NON-LACTOSE FERMENTERS 1. Lactose fermenters appear pink in colour due to the change in colour of neutral red indicator produced as a result of acid formation by these organisms. Ex.: E. Coli, Klebsiella, Citrobacter. 2. Non-lactose fermenter develop paler almost colouless colonies as they do not attack lactose. Ex.: Pseudomonas, Proteus. Salmonella and Vibrios (Fig. 60.4).

Figure 60.3: Clostridium tetani with terminal spores

MAcCONKEY’S AGAR (FIG. 60.4) 1. It is differential or indicator medium, used for the differentiation of lactose fermenters from non-lactose fermenters.

Figure 60.4: MacConkey’s agar with LF colonies

Spotters 259 2. It consists of 2% peptone, lactose, neutral red indicator, and sodium taurocholate. 3. Lactose fermenters appear pink in colour and non-lactose fermenters almost colourless or paler. CLOSTRIDIUM TETANI WITH TERMINAL SPORES 1. Gram positive bacilli. 2. Terminal spores give drum stick appearance to bacilli. 3. Spores do not take up the stain. SUGAR MEDIA TO TEST FOR CARBOHYDRATE FERMENTATION Sugar media are used to test the biochemical reactions of various bacteria. These media consist of 1% desired sugar in peptone water, along with appropriate indicator like, Andrade’s indicator. A small tube (DURHAM’S TUBE) is kept inverted in the sugar tube to detect gas production. Development of pink colour in the medium indicates acid production. Bacteria producing Acid only: a. Staphylococci b. Salmonella Typhi Bacteria producing Acid and Gas: a. Escherichia Coli b. Klebsiella. Non-fermenting bacteria: a. Moraxella b. Alkaligenes GRAM NEGATIVE BACILLI 1. Pinkish, rod-shaped bacteria. 2. Short and straight with a discrete arrangement. Ex: a. Klebsiella b. E.coli c . Salmonella, etc.

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RABBIT Uses 1. For the production of immune sera such as agglutinating and neutralising sera by intravenous inoculation of antigens. 2. In Pyrogen tests. 3. For the maintenance of pathogenic strains of Treponema pallidum intheirtesticles. 4. In Paul’s test, i.e. Production of keratitis by inoculation of Small pox virus into the scarified rabbit cornea. 5. Rabbit blood is used in the preparation of blood agar. MICE Uses 1. Used extensively for isolation of Toxoplasma gondii. 2. Suckling mice are used for isolation of Herpes simplex, Enteric viruses and Arbo-viruses. 3. To test the toxigencity of teatanus becci.24 day-old cooked deat culture is inoculated into the root of the tail of a mouse. If the strain is toxigenic, there will be stiffness of the tall. GUINEA PIG Uses 1. In the diagnosis of Weil’s disease for the isolation of Leptospira icterohaemorrhagiae. 2. For the diagnosis of murine typhus, i.e. to demonstrate NeillMosser reacton. 3. Guinea pig serum is used extensively in complement fixation test as a source of complement.. 4. To determine the virulence of diphtheria bacilli by intravenous inoculation. VDRL (VENEREAL DISEASE RESEARCH LABORATORY) TEST SLIDE 1. This is a paraffin-ringed slide used to perform VDRL test. 0.05 ml of inactivated patient’s serum is taken in each ring and a drop of

Spotters 261 VDRL antigen is added. The slide is rotated at 120 rotations per minute in a VDRL rotator for 4 minutes. Then it is examined under the microscope. Formation of Clumps indicates that the serum is reactive. 2. VDRL test is a rapid, serological slide flocculation test for syphilis. WIDAL TEST Uses 1. An agglutination test used in the diagnosis of enteric fever caused by Salmonella spp. 2. Antigens used : a. ‘O’ or somatic antigen. b. ‘H’ or flagellar antigen 3. The diagnostic titre is 1: 160 and more. 4. Positive agglutination is observed in the IInd week of infection. 5. Demonstration of a rise in titre of antibodies, by testing two or more serum samples, is more meaningful. ANTIBIOTIC SENSITIVITY PLATE 1. This is a method for testing the sensitivity of different bacteria to various antibiotics. 2. It consists of inoculated nutrient agar medium containing different filter paper discs impregnated with known concentrations of various antibiotics. 3. After the incubation at 37oC for 18-24 hours. Inhibition zones are observed around those antibiotic discs to which the bacteria are sensitive No zones are noticed if the bacteria are resistant to antibiotics. CORYNEBACTERIUM DIPHTHERIAE 1. Gram positive bacilli. 2. The bacilli are arranged in characteristic Chinese Letter or Cuneiform arrangement, resembling the letters V, N, L, T, etc. 3. Non-sporing non-capsulated and pleomorphic.

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UREASE POSITIVE AND UREASE NEGATIVE REACTIONS The production of the enzyme urease by bacteria can be tested by growing the organism in a medium containing urea. Christensons’s Medium is used for this purpose. Urease decomposes urea resulting in the formation of ammonia, the production of which is detected by change in colour of the indicator, phenol red present in the medium. Deep pink colour indicates the production of urease. Urease negative bacteria do not produce any colour change in the medium. 1. Urease positive bacteria a. Proteus species b. Klebsiella pneumoniae 2. Urease negative bacteria a. Escherichia coli b. Salmonella DERMATOPHYTE ON SABOURAOUD’S AGAR 1. Colonies are fluffy, cottony with white mycelia. 2. Reverse side of the colony is deep wine or brown in colour. 3. They are the common cause of tinea pedis, tinea corporis, tinea unguim, tinea barbae and tinea capitis. CANDIDA IN GRAM’S SMEAR 1. Gram positive, oval yeast cells. 2. Characteristic buds and pseudomycelia are seen. 3. Non-capsulate. ASPERGILLUS 1. Responsible for aspergillosis. 2. It is a fungus having septate mycelia. 3. It has long unbranched condiiophore. The tip of the condidiophore is vesicle like and possesses bottle shaped sterigmata giving a brush-like appearance. 4. Grows well on Sabouraud’s agar.

Spotters 263 ASPERGILLUS ON SABOURAUD’S AGAR 1. The common saprophytic, contaminant fungi, occasionally causing opportunistic infection. 2. The colonies grow more slowly with characteristic blackish pigmentation of conidial heads, an outstanding feature of Aspergillus niger species. MYCOBACTERIUM TUBERCULOSIS ON LOWENSTEINJENSEN MEDIUM 1. EUGONIC (luxuriant) growth observed. 2. The colonies are dry, raised rough, tough and buff with wrinkled surface un-like that of M.bovis, where the colonies are moist, flat, smooth and white. MYCOBACTERIUM TUBERCULOSIS IN ACID-FAST SMEAR (FIG. 13.1) 1. Zeihl-Neelsen’s smear showing reddish-Pink acid-fast bacilli which are found singly, in pairs and small groups. 2. Non-sporing and non-capsulated. LOWENSTEIN-JENSEN MEDIUM 1. This is a special medium used for the cultivation of human type of the tubercle bacilli. 2. It consists of beaten egg. Malachite green solution, mineral salt solution and glycerol. 3. This medium is sterilised by inspissation OVUM ASCARIS LUMBRICOIDES (ROUNDWORM) (FIG. 54.9) Fertilised

Unfertilised

1. Round or oval

Narrower and longer

2. Bile stained.

Bile stained.

3. Surrounded by Contains thinner shell and very little of a. Thick, smooth, transparent shell. albuminous coat. b. Albuminous coat, thrown into folds

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4. Contains large unsegmented ovum

Contains highly refractile granules.

5. Floats in saturated common salt solution

Does not float in saturated common salt solution.

OVUM OF ANCYLOSTOMA DUODENALE (HOOKWORM) (FIG. 54.6) 1. 2. 3. 4. 5.

Oval or elliptical. Not bile-stained. Surrounded by transparent hyaline shell-membrane. Contains segmented ovum with 4 blastomeres. Floats in saturate common salt solution.

Figure 60.5: Hydatid cyst

OVUM OF TRICHURIS TRICHURA (WHIP WORM) (FIG. 54.7) 1. 2. 3. 4.

Barrel shaped with mucous plug at each pole. Has a double shell and bile stained. Contains an unsegmented ovum. Floats in saturated common salt solution.

Spotters 265 MALARIAL PARASITE (GAMETOCYTE) 1. End products of asexual cycle capable of sexual function in the body of the definitive host (Mosquito). 2. Male cells are called microgametocytes and female cells as macrogametocytes. 3. They are spherical or rounded in P. vivax, P. Malariae and P. Ovale and cresent shaped in P. falciparum. 4. In stained smear, cytoplasm in bluish with red nucleus which is diffuse and lateral in make and compact and peripheral in female. MICROFILARIA 1. 2. 3. 4. 5.

Larva of filarial worm. Causes filariasis. It has got a blunt head and a pointed tail. Body is covered with a hyaline sheath. Somatic cells extend from the head to the tail end.

MALARIAL PARASITE (TROPHOZOITE) 1. First stage in the erythrocytic schizogony of malarial parasite. 2. In stained smear it has signet ring appearance; cytoplasm is bluish and nucleus appears red. 3. Trophozite after a period of growth gives rise to schizont. HOOKWORM (ANCYLOSTOMA DUODENALE) (FIG. 54.6) 1. Small, greyish white and cylindrical. 2. Anterior end is bent dorsally giving hook-like appearance. 3. Causes ancylostomiasis characterised chiefly by anaemia. ROUND WORM (ASCARIS LUMBRICOIDES) (FIG. 54.9) 1. 2. 3. 4.

Longest intestinal nematode. Resembles earthworm. Causes ascariasis (round worm infection). Round or cylindrical.

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5. Male:

. i ii. Female: . i ii.

Tail end curved ventrally. Vulvar waist absent. Tail end straight. Vulvar waist present at the junction of anterior 1/3, and the middle 1/3, of the body.

WHIP WORM (TRICHURIS TRICHURA) (FIG. 54.7) 1. Anterior 3/5 is thin and posterior 2/5 is thick giving a whip like appearance. 2. In male, caudal extremity is ventrally coiled. 3. Found in vermiform appendix. 4. IT causes trichuriasis. HYDATID CYST (FIG. 60.5) 1. Larva of Echinococcus granulosus is responsible for the development of hydatid cyst. 2. This is found in the intermediate host (man) 3. The cyst wall consists of 2 layers of ectocyst (cuticular layer) and endocyst (germinal layer). 4. The ectocyst has the appearance of the white of hard boiled egg and curl when cut. 5. Contains hydatid fluid which is responsible for the anaphylactic symptoms in hydatid disease. TAPE WORM (FIG. 60.6) 1. Flat, long, ribbon like worm with segmented body. 2. Two species Taenia solium (pork tape worm) and Taenia saginata (beef tape worm). 3. Body is divided into scolex, neck and strobila.

Figure 60.6: Beef tapeworm

Index ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○ ○

A Accidents and spills 7 Accidents in laboratory 7 management 8 Acid-fast staining 41 Albert’s stain 56 method 57 observation 57 requirement 57 Anaerobic cultivation 84 classifications 84 obligate aerobes 84 obligate anaerobes 84 facultative anaerobes 84 microaerophilic organisms 85 McIntosh and Filde’s anaerobic jar 87 source 85 methods of creating anaerobiosis 85 Anti-streptolysin 150 principle 150 procedure 150 requirements 150

B Babes-Ernst granules 56 Bacterial motility 40 Bacterial pathogens and diseases 88 Bacteriological examination of water 233 Eijkman test 238 method of analysis 236

procedure 236 reporting 238 standards 238 water sampling 233 dug well 235 reservoir 235 tap or pump outlet 235 Bacteriological media 69 preparation and checking 70 blood agar 74 blood tellurite agar 75 buffered glycerol saline 75 chocolate agar 74 glucose broth 71 Loeffler serum medium 75 nutrient agar 71 nutrient broth 71 types 69 differential and selective media 70 enriched media 69 ordinary culture media 69 Bile solubility test 133 principle 133 procedure 134 requirements 133 result 134 Biochemical tests 106 Biohazard waste management 33 Biological safety cabinets 3 Brownian movement 40 Brucella agglutination test 148 materials required 148 principle 148 procedure 148 Burial 36

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C Capillary tube 40 Catalase test 110 method 110 principle 110 requirements 110 result 111 Cell wall theory 52 Citrate utilisation test 124 method 125 principle 124 requirements 124 result 125 Cleaning of glassware 62 chromic acid cleaning 64 cleaning of pipettes 66 new glass wares 62 used glasswares 62 Clinical materials for microbiological investigations 95 blood for serological tests 101 materials for mycological investigations 102 hairs 102 materials from systemic fungal infections 102 nail 102 skin scrapings 102 swabs from mouth and vagina 102 materials for sterility tests 105 specimen for bacteriological investigations 96 blood for culture 96 cervical and vaginal swabs 100 conjunctival swab 100 CSF 99 ear swab 99 nasal, nasopharyngeal and oral swabs 99 pus 100 rectal swab 99

serous fluid 100 sputum 99 stool samples 99 throat swab 99 urethral swab and prostatic fluid 100 urine 96 specimens for acid-fast bacilli 100 CSF 101 faeces 101 gastric lavage 101 laryngeal swab 101 pleural and peritoneal fluids 101 pus from cold abscess 101 sputum 100 tissues 101 tissues smear for lepra bacilli 101 urine for tubercle bacilli 100 specimens for anaerobic culture 101 blood 101 prostatic fluid 101 pus 101 serous fluid 101 urine 101 specimens for parasitological examination 102 blood for microfilaria 102 faeces 102 specimens for virological investigation 103 Coagulase test 135 principle 135 procedure 136 result 136 Compound microscope 18 illuminating parts 21 electric bulb 22 filter holder 21 iris diaphragm 21 mirror 21 sub-stage condenser 21

Index 269 magnifying parts 20 eyepieces 20 objectives 20 mechanical parts 18 adjustment knobs 19 base 18 limb 19 revolving nose piece 20 stage 19 CRP screen latex agglutination slide test 152 interpretation of results 154 qualitative estimation 153, 154 semiquantitative estimation 154 principle 152 procedure 153 quality control 154 requirements 153 specimen 153 Culture of fungi 179 culture media 179 Cytoplasmic theory 52 Czapek’s dox agar 180

D Diseases of 103, 104 central nervous system 103 eye 104 respiratory tract 104 skin 104 Disposal options 35 Durhams’s tube 259

E ELISA 165, 169, 170 antibody detection 165 procedure 167 regents 166 antigen detection 169 procedure 169 reagents 169

types 170 competitive 171 inhibition 171 stick 171 Experimental animals 172 guinea pigs 173 mice 174 rabbits 172 rats 174

F First aid box 10

G Germ tube test 195 method 196 observation 196 requirements 195 Gram’s stain 41, 49 bacilli 51 observation 51 procedure 49 requirement 41, 49

H Hand washing 12 method 12 procedures 13 Hanging drop preparation 37 observation 39 procedure 38 requirements 37 Heat fixed smear 44 Hugh and Leifson’s test 108 method 109 requirements 108 Hydrogen sulphide production test 120 list of media used for H2S detection 120 method 1 120 method 2 121

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principle 120 result 121 Hypodermic syringes and needles 4

I Identification of bacteria 106 Identification of fungal isolates 184 methods 184 macroscopic examination 184 microscopic examination 185 morphology of identification 185 aspergillus 191 Candida albicans 186 Cryptococcus neoformans 185 epidermophyton floccosum 190 microsporum canis 189 mucor 194 pencillium 193 rhizopus 193 trichophyton rubrum 187 Indian ink staining 47 observation 47 staining procedure 47 Indole test 130 method 132 principle 130 requirements 130 result 132 Inoculation of culture media 77 aseptic technique 77 incubation of cultures 82 inoculation of fluid media 82 inoculation of media in petri dishes 79 inoculation of slopes 81 inoculation of stab media 82 labelling of inoculated media 82

temperature of incubation 83 wire loop 79

L Laboratory access 5 Laboratory biosafety levels 1 Laboratory clathing 7 Laboratory facilities in BSL-2 2 Lactophenol cotton blue mount 177 method 178 requirements 177 Leishman’s stain 59 method 59 observation 61 requirements 59

M Malaria 201 lab diagnosis 201 blood smear 201 examination of stained slides 205 examination of thick film 206 examination of thin film 206 Giemsa staining 205 malaria parasite 206 thick smear 202 thin smear 205 Medical entomology 220 cyclops 231 housefly 226 diseases transmitted 226 morphology 227 itch mite 229 morphology 230 louse 228 morphology 229 mosquitoes 221 aedes mosquitoes 223 anopheles mosquitoes 222 culex mosquitoes 223 mansonia mosquitoes 224

Index 271 rat flea 227 diseases transmitted 227 sandyfly 225 morphology 225 ticks 230 hard ticks 230 soft ticks 231 tsetse flies 226 morphology 226 Methyl red test 128 method 129 principle 128 requirement 128 result 129 Methylene blue staining 45 requirements 45 staining method 45 uses 46 Microbiology of milk 239 common test 239 mehtylene blue reduction test 239 phosphatase test 240 test for colifrom bacteria 239 turbidity test 240 viable count 239 Micrometry 24 calibration of eyepiece micrometer 25 measuring object 26 method 24 requirement 24 eyepiece micrometer 24 stage micrometer 24 Microscope 17 types 17 compound 17 dark ground 17 electron 17 fluorescent 17 inverted 17 phase contrast 17 polarizing 17 simple 17 ultraviolet 17

Modified Kirby-Bauer method 140 quality assurance 144 requirements 140 antibiotic discs 140 Mueller-Hinton agar 140 swabs 141 turbidity standard 141 results 143 Motility of anaerobic bacteria 40

N Neisser’s staining 56 Nitrate reduction test 118 method 119 principle 118 requirement 118 result 119

O Oxidase test 112 methods 112 dry filter paper method 113 plate methods 113 wet filter paper method 113 principle 112 requirements 112

P Parasitological examination of faeces 210 collection of faecal sample 210 macroscopic examination 211 microscopic examination 212 concentration techniques 212 procedure 212 saturated salt flotation technique 215 transportation of samples 211 pH in microbiology 67 comparater and capillator methods 68 methods used for measurement 67 pH indicator papers 68

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Pipetting 2 Plastic pasteur pipettes 65 Potassium hydroxide wet mount 175 method 175 observation 176 requirements 175 slide KOH mount 176 Preparation and fixation of smears 41 requirements 41 procedure 42 drying of smears 42 fixation of smears 43 labeling of slides 44 making of smears 42 Preparation of glass pasteur pipette 65 Protozoan parasites 37

Q Qualitative serum test 158 Quantitative serum test 158

R Riddle’s method 182 method 183 observation 183 requirements 182 Ryle’s tube 101

S Sabouraud’s dextrose agar 180 Simmons’s citrate media 125 Sperm motility 37 Spotters 253 agar agar 255 antibiotic sensitivity plate 261 aspergillus 262 aspergillus on Sabouraud’s agar 263 blood agar 255 candida in Gram’s smear 262 candle jar 253 centrifuge 254

corynebacterium diphtheriae 261 dermatophyte on Sabouraoud’s agar 262 gelatin 255 glucose broth 254 gram negative bacilli 259 gram-positive bacilli 257 guinea pig 260 haemolysis on blood agar 256 hookworm 265 hydatid cyst 266 Lowenstein-Jensen medium 263 MacConkey’s agar 258 malarial parasite 265 McIntosh and Filde’s jar 257 mice 260 microfilaria 265 Neisseria gonorrhoeae 257 nutrient agar 255 ovum of ancylostoma duodenale 264 ovum of Trichuris trichura 264 platinum wire loop 253 rabbit 260 Robertson’s cooked meat medium 257 round worm 265 Sabouraud’s glucose agar 255 seitzfilter 254 sugar media to test for carbohydrate fermentation 259 tape worm 266 testing of autoclaves 253 thioglycollate medium 257 throat swab 254 urease positive and urease negative reactions 262 VDRL 260 whip worm 266 widal test 261 Sterilization 27 autoclave 27

Index 273 gravity displacement 30 pressure-cooker type 28 boiling 32 disinfection 32 heat 27 hot air oven 31 operating instructions 31 Sugar fermentation test 115 method 116 principle 115 requirements 115 result 117

T Treponema pallidum haemagglutination assay 159 observation 160 principle 159 quantitative test 160 results 160 Tuberculosis 241 lab diagnosis 241 biosafety 246 collection of sputum sample 242 culture 244 grading of microscopy smears 243 microscopy of sputum 241 morphology 241 preparation of smear and Ziehl-Neelsen staining 242 storage and transportation of specimens 242 Tumbling motility 40

U Units 14 SI units 14 prefixes 15 Urease test 122 method 123 principle 122

requirements 122 result 123 Urinary tract infection 247 causative organisms 247 culture 250 screening procedures 250 specimen collection 248 infant and young children 249 men 248 women 249 specimen transport 249

V VDRL test 156 preparation of antigen 157 preparation of serum 157 requirements 156 Virus infections 197 diagnosis 197 embryonated egg inoculation 198 serology 198 tissue culture 198 Voges-Proskauer test 126 method 127 principle 126 requirements 126 result 127

W Widal test 162 observation 163 principle 162 procedure 163 requirement 162 result 163

Z Ziehl-Neelsen’s stain 53 requirements 53 procedure 53 observation 54