Laboratory Applications in Microbiology: A Case Study Approach [4 ed.] 9781259705229, 1259705226, 9781260418774, 1260418774, 9781260418781, 1260418782

Table of contents :
Cover
Title
Copyright
Contents
Preface
ESSENTIAL LABORATORY SKILLS
Case Study Exercise 1 Safety Considerations in the Microbiology Laboratory
CASE STUDY
Community College Acquired Infection with Salmonella Typhimurium—United States, 2017
Laboratory Contracted Plague—Chicago, Illinois, 2009
Laboratory Researcher Dies after Suffering Burns—Los Angeles, California, 2009–2018
Case Study Exercise 2 Microscopy and Measurement of Microscopic Specimens
CASE STUDY
Correspondence between Anton van Leeuwenhoek and the Royal Society of London for the Improvement of Natural Knowledge
SURVEY OF MICROORGANISMS
Case Study Exercise 3 A Survey of Protists
CASE STUDY
Primary Amebic Encephalitis—Louisiana, 2011
There's More to the Story … Phytoplankton Monitoring Network
Case Study Exercise 4 A Survey of Fungi
CASE STUDY
Histoplasmosis Outbreak among Day Camp Attendees—Nebraska, June 2012
Case Study Exercise 5 A Survey of Parasitic Worms
CASE STUDY
Trichinellosis Caused by Consumption of Wild Boar Meat—Illinois, 2013
Anisakiasis after a Meal of Sushi—Portugal 2017
Case Study Exercise 6 Ubiquity of Microorganisms
CASE STUDY
Engineering Infection Control Through Facility Design Outbreak of Multidrug-Resistant Pseudomonasaeruginosa Related to Intensive-Care Room Design—Toronto, 2009
Mucormycosis of Solid Organ Transplant Recipients at an Acute Care Hospital—Pennsylvania, 2015
MANIPULATION, STAINING, AND OBSERVATION OF MICROORGANISMS
Case Study Exercise 7 Aseptic and Pure Culture Techniques
CASE STUDY
Typhoid Fever—Colorado, 2015
Case Study Exercise 8 Simple Staining, Negative Staining, and Gram Staining
CASE STUDY
Multistate Outbreak of Listeriosis Associated with Jensen Farms Cantaloupe—United States, 2011–2015
Case Study Exercise 9 Capsular Staining
CASE STUDY
Pneumococcal Sepsis after Autosplenectomy—2005
Case Study Exercise 10 Endospore Staining
CASE STUDY
Inhalation Anthrax Associated with Dried Animal Hides—London, 2008
There's More to the Story … Isolation and Identification of Bacillus
Case Study Exercise 11 Acid-Fast Staining
CASE STUDY
Tattoo-Associated Nontuberculous Mycobacterial Skin Infections — New York, 2011–2012
ENVIRONMENTAL INFLUENCES ON THE GROWTH OF MICROORGANISMS
Case Study Exercise 12 Viable Plate Count
CASE STUDY
Campylobacter jejuni Infections Associated with Raw Milk Consumption—Utah
Case Study Exercise 13 Cultivation of Anaerobes
CASE STUDY
Large Outbreak of Botulism at a Church Potluck—Ohio, 2015
Case Study Exercise 14 Temperature Effects on Bacterial Growth and Survival
CASE STUDY
Outbreak of Gastroenteritis Associated with Consumption of Oysters—Multiple States, 2004–2013
Vibrio parahaemolyticus Infection Traced to Blue Point Oysters—Connecticut
Case Study Exercise 15 pH and Microbial Growth
CASE STUDY
Outbreak of Food borne Associated Botulism Associated with Improperly Jarred Pesto—Ohio and California
Case Study Exercise 16 Effects of Osmotic Pressure on Bacterial Growth
CASE STUDY
Vibrio vulnificus Septic Shock Due to a Contaminated Tattoo—Texas, 2017
CONTROL OF MICROBIAL GROWTH
Case Study Exercise 17 Lethal Effects of Ultraviolet Light
CASE STUDY
Gastrointestinal Outbreak Traced to Interactive Fountain—New York, March 2006
There's More to the Story … Effects of Environmental UV Exposure on Bacterial Populations
Case Study Exercise 18 Evaluation of Disinfectants
CASE STUDY
Outbreak of Skin Infections on High School Wrestling Team—Michigan, 2013
There's More to the Story … Disk Diffusion Testing of Antimicrobial Chemicals
Case Study Exercise 19 Effectiveness of Hand Scrubbing
CASE STUDY
Puerperal Fever—Vienna, Austria, 1847
There's More to the Story … Tracking Hand Hygeine
Case Study Exercise 20 Antimicrobic Sensitivity Testing: Kirby-Bauer, Tube Dilution, and ETEST® Methods
CASE STUDY
Pan-Resistant New Delhi Metallo-Beta-Lactamase-Producing Klebsiella pneumoniae—Washoe County, Nevada, 2016
There's More to the Story … Isolation of Antibiotic Producing Actinomyces
EPIDEMIOLOGY
Case Study Exercise 21 Simulated Epidemic
CASE STUDY
Measles Outbreak—Minnesota, April–May 2017
Case Study Exercise 22 Morbidity and Mortality Weekly Report
CASE STUDY
Social Media Used to Track Outbreaks of Food Poisoning—Chicago, New York, Las Vegas, 2016
There's More to the Story … Using Social Media to Track Disease
MICROBIAL GENETICS
Case Study Exercise 23 Bacterial Transformation
CASE STUDY
Vancomycin-Resistant Staphylococcus aureus—Delaware, 2015
Case Study Exercise 24 The Ames Test
CASE STUDY
Zika Virus Update—United States—2017
APPLIED MICROBIOLOGY
Case Study Exercise 25 DNA Extraction from Bacterial Cells
CASE STUDY
IKEA Pulls Meatballs from Shelves after Horsemeat Found—2013
Case Study Exercise 26 DNA Profiling
CASE STUDY
Multistate Outbreak of Salmonella Infections Associated with Peanut Butter and Peanut Butter-Containing Products—United States, 2008–2015
Case Study Exercise 27 Blood Typing
CASE STUDY
Transfusion Reaction Leads to Death Due to ABO Incompatibility—Florida, 2008
Case Study Exercise 28 Rapid Identification of Staphylococcus aureus Using Latex Agglutination Testing
CASE STUDY
Invasive Staphylococcus aureus Infections Associated with Pain Injections and Reuse of Single-Dose Vials—Arizona, 2012
Case Study Exercise 29 Slide Agglutination
CASE STUDY
Salmonella Outbreak at a Preschool—Getafe, Spain, 2013
Case Study Exercise 30 Enzyme-Linked Immunosorbent Assay (ELISA)
CASE STUDY
Lymphocytic Choriomeningitis Virus Meningoencephalitis from a Household Rodent Infestation—Minnesota, 2015
Case Study Exercise 31 Biofilm Culture and Examination
CASE STUDY
Mycobacterium abscessus Infections Among Patients of a Pediatric Dentistry Practice—Georgia, 2015, Anaheim, 2017
Case Study Exercise 32 Measures of Water Quality: Most Probable Number Procedure
CASE STUDY
Boil Water Notice—Walker County, Georgia, 2017
Case Study Exercise 33 Measures of Water Quality: Membrane Filtration Method
CASE STUDY
Contamination of AIrcraft Drinking Water—HongKong, 2015
There's More to the Story … Identification of Coliforms
Case Study Exercise 34 Measures of Milk Quality: Methylene Blue Reductase Test
CASE STUDY
Lawmakers Drink Raw Milk to Celebrate Its Legality, Become Immediately Sick—West Virginia, 2016
There's More to the Story … The Science of Yogurt
Case Study Exercise 35 Bacterial Counts of Food
CASE STUDY
Food Poisoning, Several Locations—2012–2015
MEDICAL MICROBIOLOGY
Case Study Exercise 36 Isolation and Identification of Staphylococci
CASE STUDY
Tight End Retires after Staph Infection—New York, 2016
Case Study Exercise 37 Isolation and Identification of Streptococci
CASE STUDY
Late-Onset Infant Group B Streptococcus Infection Associated with Maternal Consumption of Dehydrated Placenta—Oregon, 2016
Case Study Exercise 38 Epidemiology of Gastrointestinal Illness: Differentiation of Enterobacteriaceae
CASE STUDY
Outbreaks of E. coli, Shigella, and Salmonella—United States, 2016–2017
Case Study Exercise 39 Differential White Blood Cell Count
CASE STUDY
Two Outbreaks of Trichinellosis Linked to Consumption of Walrus Meat—Alaska, 2016–2017
IDENTIFICATION OF UNKNOWN BACTERIA
Case Study Exercise 40 Identification of Bacterial Unknowns
CASE STUDY
Respiratory Disease Strikes Legionnaires' Convention—Philadelphia, Pennsylvania, 1976
LABORATORY TECHNIQUES, REAGENTS, AND ASSAYS
Analysis of Bacterial Cultures Based on Morphological Characteristics
Exercise 41 Colony Morphology
Exercise 42 Growth in Solid and Liquid Media
Staining Techniques Used for the Microscopic Examination of Bacteria
Exercise 43 Motility Methods: Wet Mount and Hanging Drop
Exercise 44 Flagella Stain
Techniques for Inoculation of Media
Exercise 45 Streak-Plate Isolation
Exercise 46 Loop Dilution
Exercise 47 Spread-Plate
Commonly Used Differential and Selective Media
Exercise 48 Fluid Thioglycollate Medium
Exercise 49 CHROMagar Orientation Medium
Exercise 50 Mannitol Salt Agar
Exercise 51 MacConkey Agar
Exercise 52 Desoxycholate Agar
Exercise 53 Endo Agar
Exercise 54 Eosin Methylene Blue Agar
Exercise 55 Hektoen Enteric Agar
Exercise 56 Xylose Lysine Desoxycholate Agar
Exercise 57 Blood Agar
Exercise 58 Motility Medium
Exercise 59 SIM Medium
Exercise 60 Kligler's Iron Agar
Exercise 61 Triple Sugar Iron Agar
Exercise 62 Lysine Iron Agar
Exercise 63 Litmus Milk
Commonly Used Biochemical Tests
Exercise 64 Oxidation-Fermentation Test
Exercise 65 Phenol Red Carbohydrate Broth
Exercise 66 Purple Carbohydrate Broth
Exercise 67 Methyl Red and Voges-Proskauer Tests
Exercise 68 Catalase Test
Exercise 69 Oxidase Test
Exercise 70 Nitrate Reduction Test
Exercise 71 Coagulase Test
Exercise 72 Citrate Test
Exercise 73 Malonate Test
Exercise 74 Amino Acid Decarboxylation Test
Exercise 75 Phenylalanine Deaminase Test
Exercise 76 Bile Esculin Test
Exercise 77 Starch Hydrolysis
Exercise 78 ONPG Test
Exercise 79 Urease Test
Exercise 80 Casease Test
Exercise 81 Gelatinase Test
Exercise 82 DNase Test
Exercise 83 Lipase Test
Exercise 84 CAMP Test
Exercise 85 PYR Test
Commercial Identification Systems
Exercise 86 API® 20E System
Exercise 87 EnteroPluri-Test System
Antimicrobial Susceptibility Tests
Exercise 88 Antibiotic Disc Sensitivity Tests
Exercise 89 ß-Lactamase Test
Quantitative Techniques
Exercise 90 Viable Plate Count
Exercise 91 Direct Cell Count
Appendix A: Spectrophotometric Determination of Bacterial Growth: Use of the Spectrophotometer
Appendix B: Use of Pipettes in the Laboratory
Appendix C: Preparation of Culture Media
Appendix D: Media, Reagents, and Stain Formulas
Glossary
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Citation preview

Laboratory Applications in Microbiology A CASE STUDY APPROACH

Fourth Edition

Barry Chess

Pasadena City College

LABORATORY APPLICATIONS IN MICROBIOLOGY: A CASE STUDY APPROACH, FOURTH EDITION Published by McGraw-Hill Education, 2 Penn Plaza, New York, NY 10121. Copyright ©2020 by McGraw-Hill Education. All rights reserved. Printed in the United States of America. Previous editions ©2015, 2012, and 2009. No part of this publication may be reproduced or distributed in any form or by any means, or stored in a database or retrieval system, without the prior written consent of McGraw-Hill Education, including, but not limited to, in any network or other electronic storage or transmission, or broadcast for distance learning. Some ancillaries, including electronic and print components, may not be available to customers outside the United States. This book is printed onacid-free paper. 1 2 3 4 5 6 7 8 9 LMN 21 20 19 18 ISBN 978-1-259-70522-9 (bound edition) MHID 1-259-70522-6 (bound edition) ISBN 978-1-260-41877-4 (loose-leaf edition) MHID 1-260-41877-4 (loose-leaf edition) Portfolio Manager: Marija Magner Product Developers: Rose Koos, Lora Neyens, and Darlene M. Schueller Marketing Manager: Valerie L. Kramer Content Project Managers: Becca Gill Buyer: Sue Culbertson Design: Matt Backhaus Content Licensing Specialists: Lorraine Buczek Cover Image: ©CDC/James Gathany (CDC Researcher); ©CDC/National Institute of Allergy and Infectious Diseases (colorized SEM); ©McGraw-Hill Education, Lisa Burgess, photographer (pneumoniae capsule); ©Courtesy of the CDC (pneumoniae growing) Compositor: MPS Limited All credits appearing on page or at the end of the book are considered to be an extension of the copyright page. The Internet addresses listed in the text were accurate at the time of publication. The inclusion of a website does not indicate an endorsement by the authors or McGraw-Hill Education, and McGraw-Hill Education does not guarantee the accuracy of the information presented at these sites.

mheducation.com/highered

Contents Preface vii

ESSENTIAL LABORATORY SKILLS Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory  1 CASE STUDY Community College Acquired Infection with Salmonella Typhimurium—United States, 2017. Laboratory Contracted Plague—Chicago, Illinois, 2009. Laboratory Researcher Dies after Suffering Burns—Los Angeles, California, 2009–2018.

Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens  9 CASE STUDY Correspondence between Anton van Leeuwenhoek and the Royal Society of London for the Improvement of Natural Knowledge.

SURVEY OF MICROORGANISMS Case Study Exercise 3  A Survey of Protists 21 CASE STUDY Primary Amebic Encephalitis—Louisiana, 2011. There’s More to the Story . . . Phytoplankton Monitoring Network.

Case Study Exercise 4  A Survey of Fungi 43 CASE STUDY Histoplasmosis Outbreak among Day Camp Attendees— Nebraska, June 2012.

Case Study Exercise 5  A Survey of Parasitic Worms 59 CASE STUDY Trichinellosis Caused by Consumption of Wild Boar Meat—Illinois, 2013. Anisakiasis after a Meal of Sushi—Portugal 2017.

Case Study Exercise 6  Ubiquity of Microorganisms 73 CASE STUDY Engineering Infection Control Through Facility Design. Outbreak of Multidrug-Resistant Pseudomonas aeruginosa Related to Intensive-Care Room Design—Toronto, 2009. Mucormycosis of Solid Organ Transplant Recipients at an Acute Care Hospital—Pennsylvania, 2015.

MANIPULATION, STAINING, AND OBSERVATION OF MICROORGANISMS Case Study Exercise 7  Aseptic and Pure Culture Techniques 81 CASE STUDY Typhoid Fever—Colorado, 2015.

Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining 95 CASE STUDY Multistate Outbreak of Listeriosis Associated with Jensen Farms Cantaloupe—United States, 2011–2015.

Case Study Exercise 9  Capsular Staining 111 CASE STUDY Pneumococcal Sepsis after Autosplenectomy—2005.

Case Study Exercise 10  Endospore Staining 115 CASE STUDY Inhalation Anthrax Associated with Dried Animal Hides—London, 2008. There’s More to the Story . . . Isolation and Identification of Bacillus.

Case Study Exercise 11  Acid-Fast Staining 121 CASE STUDY Tattoo-Associated Nontuberculous Mycobacterial Skin Infections — New York, 2011–2012.

ENVIRONMENTAL INFLUENCES ON THE GROWTH OF MICROORGANISMS Case Study Exercise 12  Viable Plate Count 127 CASE STUDY Campylobacter jejuni Infections Associated with Raw Milk Consumption—Utah, 2014.

Case Study Exercise 13  Cultivation of Anaerobes 135 CASE STUDY Large Outbreak of Botulism at a Church Potluck—Ohio, 2015.

Case Study Exercise 14  Temperature Effects on Bacterial Growth and Survival 143 CASE STUDY Outbreak of Gastroenteritis Associated with Consumption of Oysters—Multiple States, 2004–2013. Vibrio parahaemolyticus Infection Traced to Blue Point Oysters—Connecticut, 2013.

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Contents

Case Study Exercise 15  pH and Microbial Growth 149 CASE STUDY Outbreak of Foodborne Associated Botulism Associated with Improperly Jarred Pesto—Ohio and California, 2014.

Case Study Exercise 16  Effects of Osmotic Pressure on Bacterial Growth 157 CASE STUDY Vibrio vulnificus Septic Shock Due to a Contaminated Tattoo—Texas, 2017.

CONTROL OF MICROBIAL GROWTH Case Study Exercise 17  Lethal Effects of Ultraviolet Light 165 CASE STUDY Gastrointestinal Outbreak Traced to Interactive Fountain—New York, March 2006. There’s More to the Story . . . Effects of Environmental UV Exposure on Bacterial Populations.

Case Study Exercise 18  Evaluation of Disinfectants 173 CASE STUDY Outbreak of Skin Infections on High School Wrestling Team—Michigan, 2013. There’s More to the Story . . . Disk Diffusion Testing of Antimicrobial Chemicals.

Case Study Exercise 19  Effectiveness of Hand Scrubbing 181 CASE STUDY Puerperal Fever—Vienna, Austria, 1847. There’s More to the Story . . . Tracking Hand Hygeine.

Case Study Exercise 20  Antimicrobic Sensitivity Testing: Kirby-Bauer, Tube Dilution, and ETEST® Methods 189 CASE STUDY Pan-Resistant New Delhi Metallo-Beta-Lactamase-Producing Klebsiella pneumoniae—Washoe County, Nevada, 2016. There’s More to the Story . . . Isolation of Antibiotic Producing Actinomyces.

EPIDEMIOLOGY Case Study Exercise 21  Simulated Epidemic 203 CASE STUDY Measles Outbreak—Minnesota, April–May 2017.

Case Study Exercise 22  Morbidity and Mortality Weekly Report 211 CASE STUDY Social Media Used to Track Outbreaks of Food Poisoning—Chicago, New York, Las Vegas, 2016. There’s More to the Story . . . Using Social Media to Track Disease.

MICROBIAL GENETICS Case Study Exercise 23  Bacterial Transformation 221 CASE STUDY Vancomycin-Resistant Staphylococcus aureus—Delaware, 2015.

Case Study Exercise 24  The Ames Test 233 CASE STUDY Zika Virus Update—United States—2017.

APPLIED MICROBIOLOGY Case Study Exercise 25  DNA Extraction from Bacterial Cells 239 CASE STUDY IKEA Pulls Meatballs from Shelves after Horsemeat Found—2013.

Case Study Exercise 26  DNA Profiling 245 CASE STUDY Multistate Outbreak of Salmonella Infections Associated with Peanut Butter and Peanut Butter-Containing Products—United States, 2008–2015.

Case Study Exercise 27  Blood Typing  257 CASE STUDY Transfusion Reaction Leads to Death Due to ABO Incompatibility—Florida, 2008.

Case Study Exercise 28  Rapid Identification of Staphylococcus aureus Using Latex Agglutination Testing 265 CASE STUDY Invasive Staphylococcus aureus Infections Associated with Pain Injections and Reuse of Single-Dose Vials—Arizona, 2012.

Case Study Exercise 29  Slide Agglutination 271 CASE STUDY Salmonella Outbreak at a Preschool—Getafe, Spain, 2013.

Case Study Exercise 30  Enzyme-Linked Immunosorbent Assay (ELISA) 277 CASE STUDY Lymphocytic Choriomeningitis Virus Meningoencephalitis from a Household Rodent Infestation—Minnesota, 2015.

Case Study Exercise 31  Biofilm Culture and Examination 285 CASE STUDY Mycobacterium abscessus Infections Among Patients of a Pediatric Dentistry Practice—Georgia, 2015, Anaheim, 2017.

Case Study Exercise 32  Measures of Water Quality: Most Probable Number Procedure 293 CASE STUDY Boil Water Notice—Walker County, Georgia, 2017.

Case Study Exercise 33  Measures of Water Quality: Membrane Filtration Method 303 CASE STUDY Contamination of AIrcraft Drinking Water—Hong Kong, 2015. There’s More to the Story . . . Identification of Coliforms.

Contents Case Study Exercise 34  Measures of Milk Quality: Methylene Blue Reductase Test 311 CASE STUDY Lawmakers Drink Raw Milk to Celebrate Its Legality, Become Immediately Sick—West Virginia, 2016. There’s More to the Story . . . The Science of Yogurt.

Case Study Exercise 35  Bacterial Counts of Food 317 CASE STUDY Food Poisoning, Several Locations—2012–2015.

Techniques for Inoculation of Media Exercise 45  Streak-Plate Isolation  397 Exercise 46  Loop Dilution  405 Exercise 47  Spread-Plate  411

Commonly Used Differential and Selective Media Exercise 48  Fluid Thioglycollate Medium  415 Exercise 49  CHROMagar Orientation Medium 419 Exercise 50  Mannitol Salt Agar  423

MEDICAL MICROBIOLOGY

Exercise 51     MacConkey Agar  427

Case Study Exercise 36  Isolation and Identification of Staphylococci 323 CASE STUDY

Exercise 52  Desoxycholate Agar  431

Tight End Retires after Staph Infection—New York, 2016.

Case Study Exercise 37  Isolation and Identification of Streptococci 333 CASE STUDY Late-Onset Infant Group B Streptococcus Infection Associated with Maternal Consumption of Dehydrated Placenta— Oregon, 2016.

Case Study Exercise 38  Epidemiology of Gastrointestinal Illness: Differentiation of Enterobacteriaceae  343 CASE STUDY Outbreaks of E. coli, Shigella, and Salmonella—United States, 2016–2017.

Case Study Exercise 39  Differential White Blood Cell Count 353 CASE STUDY Two Outbreaks of Trichinellosis Linked to Consumption of Walrus Meat—Alaska, 2016–2017.

IDENTIFICATION OF UNKNOWN BACTERIA Case Study Exercise 40  Identification of Bacterial Unknowns 361 CASE STUDY Respiratory Disease Strikes Legionnaires’ Convention—Philadelphia, Pennsylvania, 1976.

Exercise 53  Endo Agar  435 Exercise 54  Eosin Methylene Blue Agar  439 Exercise 55  Hektoen Enteric Agar  443 Exercise 56  Xylose Lysine Desoxycholate Agar  447 Exercise 57  Blood Agar  451 Exercise 58  Motility Medium  455 Exercise 59  SIM Medium  459 Exercise 60  Kligler’s Iron Agar  465 Exercise 61  Triple Sugar Iron Agar  469 Exercise 62  Lysine Iron Agar  473 Exercise 63  Litmus Milk  477

Commonly Used Biochemical Tests Exercise 64  Oxidation-Fermentation Test  481 Exercise 65  Phenol Red Carbohydrate Broth  485 Exercise 66  Purple Carbohydrate Broth  489 Exercise 67 Methyl Red and Voges-Proskauer Tests 493 Exercise 68  Catalase Test  497 Exercise 69  Oxidase Test  501 Exercise 70  Nitrate Reduction Test  505 Exercise 71       Coagulase Test  509 Exercise 72  Citrate Test  511 Exercise 73  Malonate Test  515 Exercise 74  Amino Acid Decarboxylation Test 519

LABORATORY TECHNIQUES, REAGENTS, AND ASSAYS Analysis of Bacterial Cultures Based on Morphological Characteristics

Exercise 75  Phenylalanine Deaminase Test  523 Exercise 76  Bile Esculin Test  527 Exercise 77    Starch Hydrolysis  531 Exercise 78  ONPG Test  535

Exercise 41      Colony Morphology  381

Exercise 79  Urease Test  539

Exercise 42  Growth in Solid and Liquid Media  385

Exercise 80  Casease Test  543

Staining Techniques Used for the Microscopic Examination of Bacteria Exercise 43 Motility Methods: Wet Mount and Hanging Drop 389 Exercise 44  Flagella Stain  393

Exercise 81    Gelatinase Test  547 Exercise 82  DNase Test  551 Exercise 83  Lipase Test  555 Exercise 84  CAMP Test  559 Exercise 85  PYR Test  563

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Contents

Commercial Identification Systems Exercise 86  API® 20E System 565

Appendix A: Spectrophotometric Determination of Bacterial Growth: Use of the Spectrophotometer  599

Exercise 87  EnteroPluri-Test System 573

Appendix B: Use of Pipettes in the Laboratory  601

Antimicrobial Susceptibility Tests Exercise 88  Antibiotic Disc Sensitivity Tests  581 Exercise 89  β-Lactamase Test  585

Quantitative Techniques Exercise 90  Viable Plate Count  589 Exercise 91     Direct Cell Count  595

Appendix C: Preparation of Culture Media  605 Appendix D: Media, Reagents, and Stain Formulas  610 Glossary  618 Index  634

Preface My students laugh at me. Why? I typically begin a class by saying “the coolest thing happened in…” and relate the latest Ebola scare, botulism outbreak, or measles epidemic, growing more animated with each news headline. Okay, so maybe “cool” isn’t the best word to describe these incidents, but I get excited when microbiology becomes a topic in the news—a flu outbreak in a small town or E. coli hitting a fast food chain. And as much as my students—and let’s face it, most people—find disease outbreaks an odd thing to get excited about, microbiologists, microbiology teachers, nurses, doctors, and health officials of all stripes perk up their ears when they hear these stories. One of the best things about living in the twenty-first century is that few of us know someone who contracted polio, died of botulism, or wasted away from tuberculosis. On the flip side, vaccination rates have fallen because no one has seen a family member suffer through measles or mumps. We have in some ways become victims of our own success. Disinfection of water, the appropriate use of antibiotics, and routine monitoring and testing of the food supply have turned much of microbiology teaching into something of a history class—“This used to happen until….” But the microbes have not gone away, and the very best way to emphasize that point is to mention the news story about a scientist who dies of the plague or a pair of roommates who contract botulism from pesto sauce they bought at the farmer’s market. As microbiologists, we recognize the story in our Twitter feed or the blurb on the radio as a living example of microbiology’s impact on our daily lives, and we share that with our students. Unfortunately, news stories do not always adhere to our syllabi, and many of these cool teaching moments go unexploited.

For Whom Is This Lab Manual Written? Written for students entering the allied health fields, Laboratory Applications in Microbiology: A Case Study Approach, is designed to use real-life examples, or case studies, as the basis for exercises in the laboratory. Throughout the past few years, the number of lecture texts utilizing case studies has grown rapidly, and for good reason—case studies work! This book is a lab manual focusing on this means of instruction, an approach particularly applicable to the microbiology laboratory. All the microbiological theory in the world means little if students cannot understand the importance of a Gram stain, antibiogram, or other laboratory procedure.

What Sets This Lab Manual Apart? This book was created to make the microbiology lab a more valuable experience by reconnecting the what and how of microbiology with the sometimes forgotten why. Although Latin names, complex media, and complicated assays will always be a part of the curriculum, the context of each exercise has been expanded so the reason for completing a specific task will be clear from the outset. Every sentence was written and each photograph chosen to accomplish this goal, and the result is a laboratory manual like nothing else in the field. You’ll notice a number of changes to the book, including:

Metacognition Metacognition is often defined as thinking about thinking. To write a lab manual and not be concerned with how students are using it to learn would be irresponsible. Many references are available that accurately describe procedures, which are great tools for the accomplished microbiologist. In contrast, this book has been designed to help create accomplished microbiologists. Every exercise has been structured from the bottom up, scaffolding knowledge so students understand the goals of an exercise, anticipate errors, acquire the skills needed for success, and eventually master the topic at hand. Each laboratory activity begins with a series of Remember, Understand, Apply questions, meant to ensure students have the requisite knowledge to begin an exercise. I think you’ll agree that knowing why a particular test is performed, what is required to complete the test, and the appearance of both a negative and positive result is exactly what we’d like a student to know before beginning an activity. At the conclusion of the exercise, the student will encounter a set of Analyze, Evaluate, Create questions, meant to reinforce their facility with the topic at hand. Asking them to interpret unfamiliar outcomes, examine results in a different context, or troubleshoot problems; these questions help students make the critical leap from apprentice to microbiologist.

Case Studies The first 40 exercises include cases taken from the scientific literature or mass media. The techniques, media, and observational tools introduced in each exercise help students solve the issues presented in the case, which drives home the relevance of microbiology vii

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and hones critical thinking skills. Evidence has shown that the use of case studies boosts learning, develops critical thinking skills, increases student retention and success, and even reduces the incidence of academic dishonesty. Simply put, students learn more, learn faster, and retain more with case studies than with traditional instruction methods. Although this seems obvious to those of us who cannot wait to share the day’s news story with our class, the results are backed up by empirical evidence. In one study focused on instructors who use cases, 97 percent reported that students who were taught with cases learned new ways to think about an issue; 95 percent reported that students took a more active part in the learning process; and 92 percent reported that students were more engaged in classes. Truly remarkable numbers.

Photographs Microbiology is certainly one of the most visual of the sciences. From the beginning, we are concerned with pink versus purple, red versus white blood cells, and how to tell the difference between the egg of a flatworm and that of a roundworm. To that end we have photographs. Hundreds of photographs that explain difficult concepts. By illustrating the answers to very basic questions: “What does a mold look like?” “Is my endospore stain positive?” or “Is that an oocyst?” students can evaluate their work with the eye of someone who has seen these things before. Exercises include photographs both of those things students are likely to see (spirogyra in a pond water sample) and those that are thankfully more rare (beta hemolytic streptococci, colonies of Bacillus anthracis). Throughout the manual, you’ll notice more than 150 new photographs. Most are completely new to the manual, while others were found in the photo atlas and have since been moved to be more accessible (because no one likes flipping pages). These new photographs were chosen because they display innovative techniques, alternative methods, or simply because each was better than the photograph it replaced. Every photograph was selected with only one goal in mind: to make microbiology more understandable to the student.

There’s More to the Story There’s More to the Story serves as a jumping off point for students who want to go the extra mile. Broadly written, these 10 addendums ask students to take the exercise they’ve just completed to the next step. After Exercise 17 (Lethal Effects of Ultraviolet Light) for example, students are encouraged to study the effects of environmental UV radiation on bacterial populations. Subsequent to an exercise on algae (Exercise 3), students have a chance to receive training and become volunteer researchers for the Phytoplankton Monitoring Network. Following an exercise on milk spoilage (Exercise 34), students can make—and study—their own fermented food. Whether used as extra credit, individual exercises, or even independent study projects, There’s More to the Story requires students to do research, generate a protocol, and prove or disprove a hypothesis; in other words, act like the scientists we are training them to be.

Tech Tips New to this edition, Tech Tips (and that is the last time you will hear about them by name, I had to call them something) are, very simply, the things we say to our class every day. They are the knowledge one accumulates after doing (or watching others do) something 10,000 times. They get neither a paragraph nor a number, just a note, if you will, inserted in the text, and are meant only to pass along an occasional quick hint.

Flip Your Classroom! It’s always good when a great idea gets a name. Hardly a new concept, “flipping” refers to students accessing basic information before coming to class, leaving more time during class for problem solving, collaborative learning, and the development of higher-order critical thinking skills. From the beginning, Laboratory Applications in Microbiology has been designed for the flipped classroom. Student learning outcomes, extensive laboratory introductions, and pre-lab questions all combine to let students begin their learning at home, maximizing the scarce time available in the lab and freeing teachers to spend more time teaching.

Changes to the Fourth Edition As time passes, the microbes we study, and the manner in which we study them, evolve in concert with subjects both microbiological (biofilms, antibiotic resistance) and technological (serological testing, DNA profiling), becoming more important to a proper understanding of microbiology. With that in mind, you’ll notice changes to the fourth edition, including four new labs (Helminths, Latex Agglutination of Staphylococci, Biofilms, and Chromogenic Agar), new organizational hierarchy for some groups of organisms, new techniques, and many, many new case studies. Once again, the only question that was asked was “Will this change result in better student understanding?” Major changes to the manual include the tips previously discussed, along with the addition of new questions to every exercise, including an entirely new set of Remember, Understand, Apply questions for each activity. A more specific list of changes includes: Exercise 1: Safety Considerations in the Microbiology Laboratory ∙∙ The lab now contains an active learning exercise to familiarize students with safety equipment in the lab. ∙∙ Two new photographs were added. ∙∙ New case study has been added concerning a researcher infected with an attenuated strain of plague due to an unknown medical condition. ∙∙ The case concerning the UCLA student who died as a result of inadequate safety training has been updated. ∙∙ A new case study on a 2017 Salmonella outbreak traced to college laboratories has been added.

Preface ix

Exercise 3: A Survey of Protists ∙∙ A completely reworked lab now presents protists as a single group in accordance with the most contemporary taxonomic theories. ∙∙ The laboratory exercise now includes more than 50 new photographs. ∙∙ Slime molds and cyanobacteria—both discussion and photographs—have been incorporated into the exercise.

Exercise 19: Effectiveness of Hand Scrubbing ∙∙ A There’s More to the Story activity, featuring the iScrub lite handwashing app to track hand hygiene, has been added. Exercise 20: Antimicrobic Sensitivity Testing: Kirby-Bauer, Tube Dilution, and ETEST® Methods ∙∙ A new case study involving Carbapenem-resistant Klebsiella pneumoniae has been added.

Exercise 4: A Survey of Fungi ∙∙ Fungi are organized into five groups according to the latest scientific understanding. ∙∙ The lab contains more than two dozen new photographs.

Exercise 21: Simulated Epidemic ∙∙ A new case study involving a measles outbreak among the Somali community in Minnesota has been added.

Exercise 5: A Survey of Parasitic Worms ∙∙ Parasitic helminths are presented as a completely new laboratory exercise. ∙∙ The exercise includes 20 new photographs and 2 new case studies centering on ingestion of parasitic worms.

Exercise 22: Morbidity and Mortality Weekly Report ∙∙ A new case study involving the use of social media to track outbreaks of food poisoning in Chicago, New York, and Las Vegas has been added.

Exercise 6: Ubiquity of Microorganisms ∙∙ The lab includes entirely new pre-lab questions, along with an expanded introduction. ∙∙ A new case study on a mucormycosis outbreak at the University of Pennsylvania Medical Center has been added. Exercise 7: Aseptic and Pure Culture Techniques ∙∙ A new case study concerning typhoid fever in Colorado has been added. Exercise 8: Simple Staining, Negative Staining, and Gram Staining ∙∙ A new staining gallery displays over a dozen photographs of stained bacterial cells. ∙∙ The case study concerning listeriosis associated with cantaloupe has been updated to reflect the latest legal ramifications of the case.

Exercise 23: Bacterial Transformation ∙∙ A new case study involving the fourteenth documented case of VRSA has been added. Exercise 24: The Ames Test ∙∙ A new case study involving Zika virus and the safety of chemicals used to repel the Aedes mosquito that serves as a vector for the virus has been added. Exercise 26: DNA Profiling ∙∙ The case study concerning Salmonella infections associated with peanuts has been updated to include the latest legal ramifications for those involved. Exercise 27: Blood Typing ∙∙ The statistics for hemolytic transfusion reactions have been updated.

Exercise 12: Viable Plate Count ∙∙ A new case on an outbreak of Campylobacter jejuni infections associated with raw milk in Utah has been added. ∙∙ A new discussion of dilution and dilution factors has been added to the lab.

Exercise 28: Rapid Identification of Staphylococcus aureus Using Latex Agglutination Testing ∙∙ An entirely new exercise focused on the use of rapid, immunologically based procedures has been added. ∙∙ A new case study concerning MRSA infections linked to reuse of single-dose drug vials is included in the exercise.

Exercise 13: Cultivation of Anaerobes ∙∙ A new case study involving botulism at a church picnic in Ohio has been added.

Exercise 29: Slide Agglutination ∙∙ A new case study concerning a Salmonella outbreak linked to environmental exposure has been added.

Exercise 15: pH and Microbial Growth ∙∙ A new case study concerning botulism from pesto sauce bought from a farmer’s market has been added.

Exercise 30: Enzyme-Linked Immunosorbent Assay (ELISA) ∙∙ A new case study involving viral meningoencephalitis linked to a rodent infestation in Minnesota has been added.

Exercise 16: Effects of Osmotic Pressure on Bacterial Growth ∙∙ A new case study involving Vibrio vulnificus infection of a recent tattoo has been added.

Exercise 31: Biofilm Culture and Examination ∙∙ A new exercise focused on the importance of biofilm formation has been added.

x

Preface

∙∙ A new case study concerning Mycobacterium abscessus Infections Among Patients of Pediatric Dentistry Practices in Georgia and California has been added to the exercise. Exercise 32: Measures of Water Quality: Most Probable Number Procedure ∙∙ A new case study concerning the appearance of E. coli in the water supply serving Walker County, Georgia, has been added. Exercise 33: Measures of Water Quality: Membrane Filtration Method ∙∙ A new case study concerning water quality on aircraft is now part of the exercise. Exercise 34: Measures of Milk Quality: Methylene Blue Reductase Test ∙∙ A new case study concerning West Virginia legislators becoming ill after consuming raw milk has been added. Exercise 35: Bacterial Counts of Food ∙∙ A new case study which follows three very different incidents of food poisoning has been added. Exercise 36: Isolation and Identification of Staphylococci ∙∙ A new case study which follows news reports of an NFL player battling a staph infection has been added. Exercise 37: Isolation and Identification of Streptococci ∙∙ A new case study centered on a late case of Group B streptococcus related to a consumption of dehydrated placenta has been added. Exercise 38: Epidemiology of Gastrointestinal Illness: Differentiation of Enterobacteriaceae ∙∙ Three new case studies, each of which examines an outbreak caused by a different species of bacteria is now part of the exercise. Exercise 39: Differential White Blood Cell Count ∙∙ A new case study concerning trichinellosis linked to consumption of walrus meat has been added. Exercise 49: CHROMagar Orientation Medium ∙∙ A new exercise covering the use of chromogenic media has been added. ∙∙ Several new photographs of bacterial growing on chromogenic agar are part of the exercise. Exercise 57: Blood Agar ∙∙ A new photograph of β-hemolysis has been added to the exercise. Exercise 80: Casease Test ∙∙ A new photograph of casein hydrolysis has been added to the exercise.

Exercise 90: Viable Plate Count ∙∙ Explanations and equations for calculating dilution factors and original cell densities have been clarified. Exercise 91: Direct Cell Count ∙∙ Explanations and equations for calculating dilution factors and original cell densities have been clarified.

Progression of Exercises Promotes Active Learning Material in each of the first 40 exercises has been carefully organized so that students develop a solid intellectual base, beginning with a particular technique, mastering it, and then applying this new knowledge to a case study. Immediately following the introductory material, pre-lab questions help students to focus on the important aspects of a technique, developing a framework for what they will need to do prior to the lab, many of which require two or three periods. Between the multiday labs, questions are posed to ensure that the students understand what they have just done, the results they should expect, and the significance of those results. Post-lab questions require applying the knowledge gained from the exercise to answer more thought-provoking questions about the techniques they have just studied. Each of the first 40 exercises concludes with a case study, a real-life situation in which the technique just mastered plays a starring role. Case study questions, generally higher-order thought questions, challenge students to apply the information they’ve learned to other situations. In a quarter of the exercises, open-ended topics for study are featured (There’s More to the Story . . .) that allow students to move beyond the everyday and become true researchers. While the first 40 exercises focus on case studies, the why of microbiology, the how of the subject has not been forgotten. The final 51 exercises serve as a thorough compendium of common microbiological methods. These exercises are presented in such a way that students will develop critical thinking skills simply by deciding on a particular course of action. All similar techniques, such as selective and differential media or biochemical tests, are grouped together, and each exercise begins with student learning objectives and a brief overview. By reviewing the overview, a student may select an appropriate test, media, or staining technique from the many available, ensuring that they have decided not only what information they need, but how to go about getting it. Written to clearly guide students while also pointing out the importance of a particular technique, this portion of the manual provides detailed, well-illustrated procedures that stand by themselves or can be used in conjunction with the case studies in the front of the book. This is particularly helpful when undertaking unknowns, as each student’s unknown culture will require a unique set of procedures for complete identification. A data sheet in exercise 40 provides a single location for students to record their test results, reinforcing the importance of record keeping in the laboratory.

Preface xi

In the workplace, allied health professionals are expected to evaluate a situation and find a solution using whatever resources are available to them. This book serves as a self-contained resource, with everything a student needs to solve a problem in the microbiology laboratory. A glossary provides definitions of all microbiological terms used in the book, a rarity in the field. Appendices contain the formula of every medium and reagent used, in addition to tutorials covering universal techniques, like the use of pipettes and spectrophotometers as well as the preparation of media. Each exercise also includes a link to applicable websites, such as the CDC homepage for each pathogenic microorganism encountered. In short, this book will help students develop the ability to solve problems.

that you have helped create a better book. In the lab at Pasadena City College, a great number of people have supplied ideas, critiques, and criticisms that have helped shape this book. Special thanks go to Jessica Igoe and Sonya Valentine, two unbelievable teachers and microbiologists from whom I learn something new each semester; John Stantzos (who never uses the phrase “it’s Greek to me,” because he knows Greek—how cool is that?); and Ray Burke. (Any really bad jokes in this book are probably Ray’s.) Of course, absolutely nothing happens in the lab without the support of Mary Timmer, laboratory technician of the gods. The second group of very smart people—who do know how to make a book—are the talented professionals at McGraw-Hill Education. Undying gratitude to Senior Product Developer, Darlene Schueller, and Senior Portfolio Manager, Marija Magner. There is no way to adequately thank anyone who has been assigned to either develop or manage me, so I will inadequately thank them. Further thanks to Marketing Manager Valerie Kramer, Content Project Manager Becca Gill (again, managing me…), Designer Matt Backhaus, and Content Licensing Specialist Lorraine Buczek. The people I’ve had a chance to work with at McGraw-Hill Education are talented professionals, and I count myself lucky to work with them. More very smart people—my wife and kids—who put up with a lot, and each week that they decide not to put me out with the trash I feel blessed. They take it in stride when my office looks like a site in need of FEMA assistance and are kind enough not to rearrange things, disrupting the experiment in chaos theory that is my desk. They know when to offer encouragement and when to offer cheesesteak. Noah, Josh, and Safura, you three mean the world to me, and without your support, I couldn’t have done this. I love you all very much.

Personalize Your Lab

Reviewers

McGraw-Hill Create™ is a self-service website that allows you to create custom course materials using McGraw-Hill Education’s comprehensive, cross-disciplinary content and digital products.

Even more smart people—as always, a talented team of microbiologists had my back. The information they provided about content, procedures, what I got right, and what I could improve upon made this a better book than I could have ever written on my own. Your feedback was very valuable.

Acknowledgments

Lubna F. Abu-Niaaj Central State University

Occasionally, one of my students asks, “how do you write a book?” While my answer usually runs toward some self-important nonsense like “Well, first you have a vision of what you want to accomplish…,” the real answer is “Find of group of very smart people who are experts at what they do.” Without the help and support of a whole bunch of people, I’d still be searching for my vision. The first thank you, as always, goes to my students, who are experts…at being students. They know—and let me know— what works and what doesn’t. I know you didn’t sign up to be test subjects for every idea that pops into my head, but there is no way I could do this without your good-natured feedback. Please know

Jennifer Bess Hillsborough Community College

Extensive Flowcharts for Bacterial Identification Exercise 40 introduces the concept of bacterial identification, using a case study recounting the recognition of Legionella pneumophila as the causative agent of Legionnaires’ disease. Within this exercise, 31 flowcharts are used to help identify bacterial unknowns commonly seen in the microbiology laboratory, a far more extensive collection than the one or two found in most manuals. This exercise also serves as an introduction to the techniques section of the manual, allowing students to quickly decide which diagnostic techniques are applicable to their particular unknown culture.

A Self-Contained Resource for the Microbiology Laboratory

Benita Brink Adams State University Timothy Cox Gulf Coast State College Marcus King United States Air Force Academy Scott Layton Cowley College

xii

Preface

Chris T. McAllister Eastern Oklahoma State College Hyun-Woo Park California Baptist University Sheela Vemu Waubonsee Community College Joe Wolf Elizabethtown Community and Technical College Joni H. Ylostalo University of Mary Hardin-Baylor And that’s how you write a book.

To the Student As an introductory student in microbiology, you may find that the reasons behind a particular exercise appear overly complex. Such is the nature of science, but the reasons should, at the very least, be apparent. The first step in closing the chasm between the scientific and the everyday is to understand, always, how each step relates to

the overall objective. It is just as important to understand why you are doing something as it is to understand what it is you are doing. If you can master both the why and the what, then your success in microbiology will be assured. This book was written with you in mind, with each feature designed to support something else. Put another way, the introductory material helps to explain the case study, the photographs and diagrams are used to clarify procedures, the glossary contains definitions of microbiological terms, and websites are provided if you would like further information on a topic. When you are using this book, please, use this book. If the meaning of a sentence is unclear, look to the accompanying figure; if a word is a mystery, use the glossary; if space is provided for a detailed drawing, give it your best shot—it will all be important soon. A well-used book becomes weathered as knowledge moves from the book to the reader, and a lab book is no different in this regard. Dog-eared pages, drawings, notes, and circled definitions are all part of learning, and the physical process of making the book yours parallels the intellectual process of making the information yours. This is as true with microbiology as it is with any other interest, job, or hobby. Take the steps to own the book, and you’ll own the information within. Good luck. Work hard and have fun.

About the Author Barry Chess has taught microbiology at Pasadena City College for more than 20 years. Prior to that, while studying at the California State University and the University of California, he conducted research into the expression of genes involved in the development of muscle and bone. At Pasadena City College, beyond his usual presence in the microbiology laboratory and lecture hall, Barry has taught majors and nonmajors biology, developed a course in human genetics, helped to found a biotechnology program on campus, and regularly supervises students completing independent research projects in the life sciences. Over the past several years, his interests have focused on innovative methods of teaching that lead to greater student success. He has written and reviewed cases for the National Center for Case Study Teaching in Science and contributed to the book Science Stories You Can Count On: 51 Case Studies with Quantitative Reasoning in Biology. Barry has presented papers and talks on the effective use of case studies in the classroom, the use of digital tools to enhance learning, and for several years served as a scientific advisor for the American Film Institute. In addition to Laboratory Applications in Microbiology, Barry is coauthor of the lecture text Foundations in Microbiology, now in its tenth edition. He is a member of the American Association for the Advancement of Science and the American Society for Microbiology. Barry was profiled in the book What Scientists Actually Do, where he was illustrated as a young girl with pigtails, about to stick a fork into an electrical outlet.

©Noah Chess

xiii

LABORATORY SAFETY GUIDELINES The microbiology lab presents a number of safety challenges, many of which are unique to this field of study. The following guidelines should be followed to help ensure your safety in the microbiology laboratory. 1.   Be realistic if you feel you shouldn’t be in the lab because of health concerns. Conditions that may leave you vulnerable to infection such as a short-term illness, being immunocompromised, taking immunosuppressant drugs, or being pregnant should be candidly discussed with your instructor. 2.   Dress appropriately for the lab. No open toed shoes or sandals. Clothing with baggy sleeves that could catch fire or hinder your movements should be avoided. 3.   Know where the safety equipment is in the lab. Note the location of the eye wash, safety shower, fire extinguisher, and first aid kit. Take a moment to learn their operation. 4.   Always wear a lab coat while in the lab. Even if you are not working yourself, another person in the lab could have an accident. This garment should only be used during lab and should remain in the laboratory. 5.   Wash your hands prior to beginning lab and just before leaving as well. Also wash when removing gloves and if you feel you may have contaminated yourself. 6.  Tie back long hair. It is both a source of contamination and a fire hazard. 7.   Nothing should go into your mouth during lab. Do not eat or drink in the lab, even if no work is being done at the time. 8.   Do not apply makeup and never handle contact lenses in the lab. 9.  Always wear gloves when handling blood or blood products. 10.  Wash with an antiseptic if your skin is exposed to microorganisms as a result of a spill. 11. Dispose of broken glass, needles, lancets, wooden applicators and any other object that could penetrate the skin, in a hard sided sharps container. Do not overfill the container and never, ever force objects into the container. 12.  Make use of fume hoods when undertaking procedures where noxious chemicals may be released during heating. 13.  In the event of a spill, notify your instructor immediately.

C A S E S T U DY E X E R C I S E

1

Safety Considerations in the Microbiology Laboratory STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Demonstrate proper primary and secondary containment procedures. 2. Explain the procedures involved in dealing with a laboratory emergency.

INTRODUCTION The microbiology laboratory presents a number of unique challenges. Not only are the normal hazards of a laboratory environment (flames, caustic chemicals, and glassware) present, so too are infectious organisms. In fact, the microbiology lab is devoted to growing and studying the very organisms that may cause us such harm! Laboratory workers are by no means immune to infection in the laboratory, and it is not an exaggeration to say that proper safety procedures can be a matter of life and death. Safe laboratory procedures revolve around containment of microorganisms. Primary containment concerns the protection of personnel and the laboratory environment from exposure to infectious microbes. Proper microbiological techniques, such as the safe transport and disposal of cultures, along with the correct use of personal safety equipment (e.g., gloves and safety glasses) go a long way toward accomplishing the goal of personal containment. Secondary containment deals with protecting the outside environment from exposure to infectious organisms and depends principally on the design of the laboratory and the availability of equipment. Most laboratory workers can do little to influence the physical aspects of the laboratory other than not disabling safety features, such as keeping open a door that should remain closed, turning off an exhaust fan, or removing a fire extinguisher. #1 Rule: No eating or drinking in the lab! After all, how would you feel if I practiced microbiology in your kitchen? The type of organisms dealt with in the laboratory will dictate the safety precautions used. Working with deadly viruses obviously requires a greater degree of vigilance than working with bacteria that are nonpathogenic. To clarify exactly what techniques and equipment should be used, microorganisms are classified into one of four biosafety levels (BSL-1 through BSL-4) based on their ease of transmission and pathogenicity. Each level has a set

of minimum standards with regard to laboratory practices, equipment, and facilities. At one end, BSL-1 organisms generally do not cause disease in a healthy person and require very few specialized techniques. BSL-4 organisms, in contrast, are easily transmitted and cause life-threatening diseases. BSL-4 laboratories are the stuff of science fiction, with full-body spacesuits, respirators, and showers upon exiting the laboratory. Table  1.1 summarizes the recommended biosafety levels for selected infectious agents. The vast majority of introductory microbiology laboratories are designed to handle BSL-1 and BSL-2 rated organisms, and the rules that apply in these laboratories have a very common sense feel about them. Your instructor may modify these rules based on college, municipal, or state regulations as well as the organisms you are likely to work with as part of your course. Adhering to these guidelines will help to ensure your safety in the microbiology lab.

Prior to the Lab ∙∙ Dress appropriately for the lab. No open-toed shoes or sandals. Clothing with baggy sleeves that could catch fire or hinder your movements should be avoided. ∙∙ Know where the safety equipment is in the lab. Note the location of the eye wash, safety shower, fire extinguisher, and first aid kit. Take a moment to learn their operation; remember, if you need to use the eyewash, you very well may not be able to see at the time.

During the Lab ∙∙ Always wear a lab coat while in the lab. Although you may not be working yourself, another person in the lab could have an accident. This garment should only be used during lab and should remain in the lab. Even discounting potential biohazards, a lab coat will protect your clothing. There is a reason many of the chemicals you will be working with are called stains. ∙∙ Wash your hands prior to beginning the lab and just before leaving as well. Also wash after removing gloves or if you feel you may have contaminated yourself. If your laboratory sink has a hands-free method of activating the flow of water (such as foot pedals), use it. ∙∙ Tie back long hair, it is both a source of contamination and a fire hazard. ∙∙ Disinfect your benchtop with amphyl, Lysol, or 10% bleach prior to beginning work and just before leaving the laboratory. If time permits, allow the disinfectant to evaporate rather than wiping the surface of your bench dry. ∙∙ Keep clutter on your bench to a minimum. Store book bags, purses, and other unneeded items where they will not consume 1

2

Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory

TABLE 1.1

Summary of biosafety levels for selected infectious agents

Health risk

Practices

Primary barriers

Secondary barriers

Organisms (Selected examples)

1

Not known to cause disease in healthy individuals.

Open bench microbiology.

None required.

Open benchtops and sinks.

Micrococcus luteus and Bacillus megaterium.

2

Can cause disease in healthy people, but organisms are easily contained.

Limited lab access and biohazard warning signs.

Gloves, lab coat, eye protection, and/or face shield as needed.

BSL-1 plus: ∙ Access to autoclave.

Escherichia coli and Staphylococcus aureus, along with most other human pathogens.

3

Can cause severe disease, especially when inhaled.

BSL-2 plus: ∙ Controlled access to lab. ∙ No unsterilized material can leave the lab. ∙ Decontamination of clothes prior to laundering.

BSL-2 plus: ∙ Biosafety cabinets used for all manipulations.

BSL-2 plus: ∙ Access to self-closing double doors. ∙ Negative pressure (air flows into lab from outside). ∙ Exhausted air not recirculated.

Mycobacterium tuberculosis, HIV, and Yersinia pestis.

4

Highly virulent microbes posing extreme risk to humans, especially when inhaled.

BSL-3 plus: ∙ Clothing must be changed before entering the lab, and personnel must shower upon exiting the lab. ∙ All material is decontaminated prior to leaving the facility.

BSL-3 plus: ∙ All procedures are conducted in complete isolation biosafety cabinets (BSCs) or in class I or II BSCs along with full-body, positivepressure suits with supplied air.

BSL-3 plus: ∙ Isolated building or lab. ∙ Isolated laboratory systems (air supply and exhaust, vacuum, and decontamination).

Lassa fever virus, Ebola virus, and Marburg virus.

BSL

precious bench space and where they will be less likely to be contaminated by an inadvertent spill. ∙∙ Put your cell phone away! Cell phones held to your mouth substantially increase the risk of accidental infection and can easily transport bacteria out of the laboratory. Switching between lab work and texting is an equally bad idea.

up away from overhead equipment or shelving, and the immediate area should be free of combustible materials such as notes or books. Prior to lighting the burner, quickly inspect the hose for holes, cracks, or leaks, and be sure it fits securely on both the gas valve and the burner.

Your cell phone has no place in the lab. Don’t rely on your phone as a calculator or timer. ∙∙ Nothing should go into your mouth while you are in the laboratory. Do not eat or drink in the lab, even if no work is being done at the time. ∙∙ Skin and eyes represent a common portal of entry for pathogens. Do not apply makeup, and never handle contact lenses in the lab. ∙∙ Organize your workplace before beginning (Figure 1.1). Store culture tubes upright in a rack, never on their side. Caps on tubes are generally not tight fitting, and liquid will leak out (even from a solid culture), leading to contamination. ∙∙ The open flame produced by a Bunsen burner presents an obvious danger in the laboratory. Burners should be set

Figure 1.1  Personal protection in the laboratory includes the use of a lab coat, gloves, and eye protection. Also note that long hair is tied back and the work area is free of clutter. ©nandyphotos/Getty Images



Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory

3

Disposal of Contaminated Materials Most material in the microbiology laboratory must be decontaminated prior to being disposed of or reused, and this is most often accomplished using an autoclave (Figure  1.2), which uses steam under high pressure to kill even the most resistant organisms. After decontamination, culture tubes, glass pipettes, and the like are washed and reused. Plastic Petri dishes melt during the decontamination process and are discarded after autoclaving along with single-use items such as tongue depressors, needles, and swabs. In general, disposal of lab materials depends on whether or not it will be reused. In any event, the contents of plates or tubes should never be touched by hand. ∙∙ Dispose of plastic Petri dishes, swabs, disposable gloves, and similar nonreusable items in a biohazard bag (Figure 1.3a). Petri dishes should be taped closed, but there is no need to remove labels or tape from items. ∙∙ Reusable supplies such as culture tubes and glass pipettes should have all labels removed before being placed in a rack or container designated for autoclaving. ∙∙ Used microscope slides should be placed in a container for autoclaving or soaked in a disinfectant solution for a minimum of 30 minutes before being discarded.

Safety Considerations ∙∙ Be realistic if you feel you shouldn’t be in lab because of health concerns. Conditions that may leave you vulnerable to infection such as a short-term illness, being immunocompromised, taking immunosuppressant drugs, or being pregnant should be candidly discussed with your instructor. ∙∙ Always wear gloves when handling blood or blood products. Bloodborne pathogens have special procedures associated

Figure 1.2  Autoclaves use steam under pressure to sterilize biohazardous materials prior to disposal. ©Barry Chess

(a)

(b)

Figure 1.3  All disposable, potentially infectious waste should be placed in a biohazard container, with needles, slides, tongue depressors, and anything else that could penetrate a plastic bag restricted to disposal in a hard-sided receptacle. The international biohazard symbol on these containers not only marks the contents for autoclaving prior to disposal but also cautions anyone in the room as to the possibly hazardous nature of the items inside the container. (a) ©McGraw-Hill Education/Tim Fuller, photographer; (b) ©McGraw-Hill Education/Sandra Mesrine, photographer

with them, and work of this type should only be done with the explicit knowledge of your instructor. ∙∙ Wash with an antiseptic if your skin is exposed to microorganisms as a result of a spill. ∙∙ Dispose of broken glass, needles, lancets, wooden applicators, and any other object that could penetrate the skin, in a hardsided sharps container (see Figure 1.3b). Do not overfill the container, and never, ever force objects into the container. ∙∙ Biosafety cabinets are sometimes used to work more safely with BSL-2 organisms (figure 1.4). BSL-3 and BSL-4 organisms have more stringent requirements.

Figure 1.4  A biosafety cabinet can be used to enhance protection when working with potential pathogens. Note that the glass shield is lowered as far as is practical and that clutter within the cabinet is kept to a minimum. The vents at the bottom of the cabinet are connected to a powerful vacuum that creates a curtain of air, further helping to protect the user from organisms being manipulated within the cabinet. ©McGraw-Hill Education/James Redfearn, photographer

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Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory

∙∙ In the event of a spill, notify your instructor immediately. Broken glass and bacterial cultures are a hazardous combination. With the instructor’s approval, cover the spill with paper towels and saturate the towels with disinfectant. After 10 minutes, carefully wipe up the spill and discard the paper towels in the biohazard container for autoclaving. Discard the broken glass in the sharps container.

PRE-LAB QUESTIONS Remember, Understand, Apply 1. A mechanism that automatically closes a door leading into a laboratory would be considered an example of a. primary containment. b. secondary containment. c. tertiary containment. d. quaternary containment. 2. The most dangerous microbiological pathogens are classified as a. BSL-1. b. BSL-2. c. BSL-3. d. BSL-4. 3. The laboratory benchtop should be cleaned at least _____ per laboratory period. a. once b. twice c. three times d. four times 4. You should refrain from eating and drinking in the lab until both you and your immediate neighbors have finished working. a. true b. false 5. Liquid cultures should be disposed of by a. pouring the contents of the tube into the sink and then washing the tube. b. discarding the tube into the nearest trash can. c. removing any labels from the outside of the tube and placing the tube in a rack for disinfecting. d. pouring disinfectant into the tube to kill the culture, then washing the tube. 6. In the laboratory, cell phones a. should be nearby in case emergency help is required. b. should be used only for texting, calculating, or the timing of reactions, so the phone is not brought close to your mouth. c. should be kept away from the bench and not used at all. d. should be disinfected at the end of the laboratory period.

7. Personal safety equipment and practices in a BSL-1 or BSL-2 laboratory would generally include all of the following except a. refraining from eating and drinking in the laboratory. b. tying back long hair. c. wearing a mask over your nose and mouth. d. wearing shoes that cover the entire foot. 8. A wooden applicator stick that had been used to sample bacteria should be disposed of a. in a wastepaper basket. b. in a biohazard bag. c. in a hard-sided biohazard container. d. in a container filled with disinfectant.

PROCEDURE 1. Identify the location of each of the following pieces of safety equipment in your laboratory. Lab coat storage

Fire extinguisher

Disinfectant

First aid kit

Safety shower

Eyewash station

Biohazardous waste disposal

Sharps container



Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory

5

REVIEW QUESTIONS Analyze, Evaluate, Create 1. Classify each of the following agents as BSL class 1, 2, 3, or 4 based on its description. Description

Severity of disease

Mode of transmission

Ebola virus

Lethal in 50%–90% of cases. No effective treatment.

Direct contact or body fluids.

Mycobacterium tuberculosis

Severe, but treatable respiratory disease.

Respiratory droplets.

Bacillus subtilis

Does not cause disease in immunocompetent persons.

Not easily transmitted.

Clostridium tetani

Can be lethal in nonprotected individuals. Vaccine provides protection.

Anaerobic sites (deep puncture wounds).

BSL level

2. Explain, as specifically as possible, how each of the following helps to enhance safety in the microbiology lab. Explain whether each is necessary in all instances. Negative air flow (i.e., air flows into the laboratory rather than out)

Gloves, safety glasses, and lab coat

Foot pedal activation of sinks

Prohibitions on eating and drinking in the lab

3. How would you properly dispose of each of the following items? A Petri dish containing a fungal culture

A glass culture tube containing a bacterial culture

A spill containing broken glass and a bacterial culture

6

Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory

CASE STUDIES The following cases illustrate the very real dangers of working in a microbiology laboratory. Study the details of each case and use your knowledge of safe laboratory practices to answer the questions which follow. Community College Acquired Infection with Salmonella Typhimurium—United States, 2017.

reported engaging in laboratory behaviors that would increase their risk of infection, including failing to wear a lab coat, inadequate handwashing, and using the same writing utensils and notebooks outside of the laboratory. If that weren’t enough, in 2017, a third multistate outbreak of Salmonella Typhimurium infections took place. Once again linked to college and university teaching laboratories, the 2017 outbreak affected 24 persons in 16 states, with 6 requiring hospitalization. Interviews with ill persons revealed many of the same lapses in safety seen in the 2011 and 2014 outbreaks.

In December 2011 three persons suffering from Salmonella Typhimurium infection were identified by the New Mexico Department of Health. Salmonella infections of this type are usually marked by diarrhea, fever, and cramps lasting four to seven days, but more severe cases require hospitalization, Scientist Dies of Laboratory Acquired Plague, and death is a distinct possibility if the bacterium enters the Chicago, 2009 bloodstream. The New Mexico Department of Health took a special interest in these three persons because one was a Dr. Malcolm Casadaban worked at the University of Chistudent and two others were children of students in microbicago studying Yersinia pestis, the bacterium responsible for ology classes held at two different community college cambubonic plague. Spread by fleas, plague has been feared puses. A complete investigation by the Centers for Disease for centuries. Epidemics in Europe and Asia were thought to Control (CDC) eventually identified 109 patients across a have killed more than 100 million people in the 1300s, shap38-state area sickened by the same bacterium. ing migrations patterns and leading to attacks on Jews, friars, Analysis by the CDC found foreigners, lepers, and other that the greatest risk factor for groups thought responsible contracting this strain of Salfor outbreaks. Known as the monella was not eating conBlack Death because of the ne­taminated hamburger, alfalfa crosis (tissue death) caused sprouts, or cantaloupe—all by the bacterium, plague conof which had been linked to tinues to cause small numbers other outbreaks—but rather of deaths around the world. being recently exposed to a Plague is a zoonotic disease, microbiology laboratory. Inadmeaning that it is most comequate adherence to safety monly encountered in animals but can be spread to humans. procedures had not only sick­ Forest rangers, campers, vetened many students and emerinarians, or anyone who ployees, but the infectious bac­ interacts with animals is at terium was taken home on increased risk of infection. pens and pencils, cell phones, Bubonic plague is often referred to as the Black Death because of Casadaban was working and backpacks, where family the necrosis caused by infection with Yersinia pestis. Source: CDC with Yersinia pestis in an at­­ members of the students were /Christina Nelson, MD, MPH then infected. The Centers for tempt to develop an effecDisease Control examined safety practices in a number of tive vaccine against the plague. The strain with which he was labs and found that students in labs where illnesses occurred working had been deliberately weakened so that it could were less likely to have biosafety training than students in labs not absorb enough iron to make functional enzymes. Even where no illnesses occurred. Because many people involved when injected into mice, the bacterium was incapable of in the outbreak were college students, the median age of causing death. those infected was 21 years. In all, 13 hospitalizations and one In mid-September, Casadaban visited his doctor, death were attributed to this completely preventable incident. complaining of “flu-like” symptoms. He was told he most Despite a renewed emphasis on laboratory safety, espelikely had contracted a virus and was directed to rest. Three cially in teaching labs, a second outbreak of the same bactedays later he returned to the doctor, very sick, and died rium was seen in 2014. Forty-one people across 13 states were 13 hours later. An autopsy revealed the supposedly innocinfected, with 36% requiring hospitalization. The median age of uous strain of Yersinia pestis in his system, but Casathose infected was 20 years, and 86% were enrolled in a bioldaban’s death remained a mystery. How could such a ogy or microbiology course. When surveyed, many ill persons weakened strain of Yersinia pestis cause death? Analysis of the



Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory

doctor’s blood finally solved the puzzle. Unbeknownst to him, Dr. Casadaban suffered from hemochromatosis, a genetic disorder in which people accumulate high levels of iron in their blood. This excess of iron allowed the usually iron-starved Yersinia pestis to assume its original virulence. Further investigation revealed that he occasionally went without gloves in the laboratory, thinking the bacterium was entirely incapable of causing infection. Dr. Ken Alexander, an infectious disease specialist and colleague of Dr. Casadaban, said that the dead researcher would have had one comment for those scientists investigating his death, “Listen guys, I’m trying to teach you something—and you better damn well learn it.”

Laboratory Researcher Dies after Suffering Burns— Los Angeles, California, 2009–2018 Sheri Sangji, a 23-year-old research assistant at UCLA, was transferring a small quantity of t-butyl lithium from one container to another when a plastic syringe came apart in her hands, splashing her with a chemical compound that ignites instantly when exposed to air. She received second and third degree burns over 43% of her body and died 18 days later. Sangji was a recent college graduate who had only been working in the lab at UCLA for a few months when the incident occurred. An investigation by the California Division of Occupational Safety and Health concluded that she had not been properly trained for the procedure she was

undertaking and did not know what to do in the event she caught fire. Sangji was not wearing a protective lab coat and was dressed in a nylon sweater described as “solid gasoline” by a lab safety expert. A previous inspection of the lab by UCLA safety personnel turned up several safety violations, which had not been corrected at the time of the accident. UCLA, as part of a plea agreement with the Los Angeles County district attorney’s office, agreed to accept responsibility for the lab’s operating conditions at the time of the fire, to follow comprehensive safety measures, and to endow a $500,000 scholarship in Sangji’s name. Patrick Harran, the professor in whose lab the accident occurred, was arrested on felony charges, the first instance of a criminal case arising from an academic laboratory accident. As part of a plea agreement, he promised to develop and teach a chemistry course for inner-city students, perform 800 hours of community service in the UCLA hospital system, and pay $10,000 to the Grossman Burn Center, where Sangji died. If the professor complies with the details of the plea agreement, the charges against him will be dismissed. Throughout the trial, UCLA and Dr. Harran portrayed Sangji as an experienced chemist who had been properly trained and chose not to wear protective gear while she worked in the lab, though others had very different opinions. Said Neal Langerman, the previously mentioned lab safety expert, concluded, “Poor training, poor technique, lack of supervision, and improper method . . . She died, didn’t she? It speaks for itself.”

CASE STUDY ANALYSIS 1. For each of the cases seen here, postulate where the breakdown in laboratory safety occurred and suggest how it could be corrected. Salmonella infection

Plague infection

Laboratory fire

7

8

Case Study Exercise 1  Safety Considerations in the Microbiology Laboratory

2. The incident of plague infection seen in this case study is interesting primarily because a medical condition of the researcher put him alone at risk of infection. Provide two or three examples of common medical or biological conditions that could make working in the microbiology lab especially hazardous. Be sure to justify your answers.

REFERENCES CDC. 2012. Investigation Update: Human Salmonella Typhimurium Infections Associated with Exposure to Clinical and Teaching Microbiology Laboratories. http://www.cdc.gov/salmonella/typhimurium-laboratory/011712/index.html Christensen, Kim. June 20, 2014. UCLA professor strikes deal in lab fire case, avoids prison. Los Angeles Times. MacKenzie, Debora. March 1, 2011. Plague scientist dies of . . . the plague. New Scientist. U.S. Department of Health and Human Services. 2009. Biosafety in Microbiological and Biomedical Laboratories (BMBL), 5th ed. www.cdc.gov/biosafety/publications/bmbl5/BMBL.pdf

C A S E S T U DY E X E R C I S E

2

Microscopy and Measurement of Microscopic Specimens STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Properly use and care for a brightfield microscope. 2. Explain the theories underlying optical microscopy.

INTRODUCTION Whether we are discussing our skin, plaque on our teeth, a glass of milk, or a swimming pool, nearly everything in our day-today existence is loaded with microorganisms. Being able to view these creatures is essential to an understanding of microbiology. It should come as no surprise that the proper use of a microscope is a central skill in microbiology, and the placement of this laboratory exercise near the beginning of this book is no accident.

LIGHT MICROSCOPE

Components of the Light Microscope The instrument most commonly seen in microbiology labs is the brightfield microscope, so named because when objects are examined, they appear as dark objects in a bright visual field. Although your microscope may differ slightly in appearance from the one seen in this exercise, in theory and practice, the same rules will apply. The components and functions of each of the parts found in a typical brightfield microscope (Figure 2.1) are outlined here. Framework  The frame of a microscope consists of the arm and base. Keep in mind when holding the microscope that these are the only two parts built to support its weight. Lamp  A light source is located in the base of a microscope. A rotating wheel or knob can be used to adjust the voltage received by the lamp, which in turn adjusts the intensity of the light. Many microscopes will also have a blue filter that can be placed over the light source to reduce the intensity of the light and increase the resolution of the microscope. Diaphragm  The diaphragm is an adjustable disc with a hole in the center. The size of the hole can be varied to allow more or less light to pass to the slide by use of a dial or lever.

Condenser  The first of three lens systems found on all microscopes, the condenser is located beneath the stage and is usually contained in the same housing as the diaphragm. The condenser collects and focuses light on the specimen being studied. Although the condenser can be raised or lowered, best results in the microbiology lab will be obtained when the condenser is kept at its highest point, just below the level of the stage. Stage  The platform that supports the slide is known as the stage. Most microscopes have a clamping device, the mechanical stage, that allows the slide to be held and moved with greater precision. Objective Lens  Three or four objective lenses, more commonly referred to as objectives, are found just above the stage. The objectives are attached to a revolving nosepiece that allows the lenses to be rotated into position. Most microscopes will have three objectives with magnifications of 10x, 45x, and 100x, designated as low power, high dry, and oil immersion, respectively. Occasionally a 4x scanning objective will also be present, but it tends to be of little use in most microbiology labs. Ocular Lens  The third set of lenses, those closest to your eyes, are the ocular lenses. In most instances, these lenses have a magnification of 10x. Binocular microscopes have two sets of lenses while monocular microscopes have only a single ocular. Binocular microscopes will also have a means of adjusting the distance between the oculars. One ocular may also have a small ring that allows the focus of that ocular to be adjusted independently of the rest of the microscope. Focus Adjustment  Two concentric focusing knobs are located on each side of the microscope. The large outer knob is the coarse focus adjustment, while the smaller, inner knob is the fine focus adjustment.

Care of the Light Microscope Microscopes are delicate instruments, and care must be taken in their use. Some general rules related to microscope care include: Transport  The microscope should always be held with two hands. One hand should grasp the microscope around the arm while the second supports the instrument from the bottom. The biggest danger in carrying a microscope with one hand is not that it will be dropped but rather that the scope will collide with the corner of a lab bench or other piece of furniture. Once at your bench, place the scope gently on the table. 9

10

Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens Interpupillary adjustment

Ocular lens (eyepiece)

Nosepiece Objective lenses (4) Mechanical stage Stage

Iris diaphragm control

Stage adjustment knobs Substage condenser

Fine focus adjustment Course focus adjustment Base with light source

Figure 2.1  Components of a typical compound microscope. ©Barry Chess

Electric Cord  Dangling electric cords are never a good idea, and more than one microscope has been pulled off a table when a hand, foot, or backpack has become entangled in the cord. Keep excess cord secured or wrapped loosely around the base of the microscope. Likewise, water and electrical devices are a poor mix. Keeping any excess cord secured or wrapped loosely around the base of your microscope decreases the chances of it becoming entangled or coming in contact with water. Protection Against Dust and Chemicals  If a dustcover is provided for your microscope, be sure to cover the instrument while staining or undertaking any other procedure that could splatter your scope with dyes or chemicals. Also be sure to cover the scope at the end of the period.

Lens Cleaning  The number one cause of unacceptable images is dirty microscope lenses. Besides impeding your ability to see, dust, oil, and other contaminants will eventually damage the lenses, which are often the most expensive part of the microscope to replace.

Lens Care The importance of lens care cannot be overstated. Dirty or scratched lenses will limit the degree of resolution achievable with your microscope, and the delicate nature of optical glass means that scrupulous attention to detail is required when attempting any cleaning. When cleaning any of the three lens systems on your microscope, follow the guidelines below as well as any labspecific instructions you may receive.



11

Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

Use only lint-free optical tissues or cotton swabs to clean lenses. Other tissues or cloths may contain microscopic grit or lint that can damage lenses. If the lenses are quite dirty, a small amount of alcohol or xylene can be used to remove oily residue. Other solvents such as acetone may be acceptable but can damage the mounting cement used on objective lenses. Check with your instructor as to which solvents are acceptable for cleaning lenses. Ocular lenses are often the recipients of thumb prints, dust, and mascara, and so are often quite dirty. The easiest method of determining if the ocular is clean is to rotate it and see if the dirt rotates as well. If the dirt rotates, clean the ocular; if it remains stationary, check the objective lenses. Occasionally dust and other debris will make their way to the inside of the ocular. If this is the case, the ocular can be removed and compressed air can be used to dislodge the particles. Whenever an ocular is removed from the microscope, a piece of lens tissue should be used to cover the ocular opening. Although removing an ocular is a straightforward procedure, consult your instructor before disassembling any part of your microscope. Objective lenses are commonly soiled with material from slides or fingers. If a properly focused image ever appears unclear or cloudy, it is safe to assume that the objective lenses are soiled. Clean the lenses using lens paper or a cotton swab moistened with a solvent acceptable in your laboratory. Use a circular motion beginning at the center of the lens and working outward. Condensers generally accumulate less dirt than oculars or objectives, but their upward-facing surfaces do tend to collect dust. An occasional wiping with a piece of lens tissue is enough to keep them clean in most cases. Prior to returning your microscope at the end of the day, it is essential to clean the immersion oil from all the objective lenses. When left on the lenses for an extended period, the oil can soften the cement holding the lenses in place, rendering the lens unusable.

(a) Brain

Retina Eye

Ocular lens

Virtual image

Objective lens

THEORY Microscopy has two related goals. The first goal is magnification, or the creation of a larger-than-life image. This image is created as light first passes through the condenser lens focusing the light on the specimen. The light refracts (Figure 2.2a), or bends, as it passes through the objective lens, creating the real image. Light from the real image is then refracted again, this time by the ocular lens. The magnification of the real image leads to production of the virtual image. The virtual image then passes into the eye where it is eventually interpreted by the brain (Figure 2.2b). The total magnification of any specimen is easily calculated by multiplying the magnifications of the objective lens and the ocular lens. Total Magnification = magnification of ocular lens

Magnification × of objective lens

For example, a 10x ocular used in combination with a 45x objective lens would provide a total magnification of 450x. Conveniently, the magnification of both ocular and objective lenses is marked on the lens (Figure 2.3).

Light rays strike specimen

Specimen Real image

Condenser lens

Light source

(b)

Figure 2.2  Refraction and the formation of a microscopic image. a) When light rays, moving here from left to right, stop travelling through air and begin to pass through the glass of the lens, they refract, or bend, eventually causing the image to be magnified. b) As light passes through the condenser, it is focused on the specimen. Light leaving the specimen is refracted by the objective lens, forming the real image. Light from the real image is refracted again, by the ocular lens, to form the virtual image that strikes the retina and is interpreted by the brain. Notice that the virtual image is reversed both left-to-right and top-tobottom with respect to the specimen. (a) ©Don Farrall/Getty Images

12

Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

Objective lens

Nonrefracted or saved light rays Specimen

Lost light rays due to refraction Air

Condenser lens

Figure 2.4  Immersion oil has the same refractive index as glass, which prevents the loss of light rays due to refraction.

Figure 2.3  The characteristics of each objective are inscribed on the barrel of the lens. In this case, the centermost objective has a magnification of 100x and a numerical aperture of 1.25. ©Barry Chess

The second goal of microscopy is resolution, which can be most easily understood as the clarity of an image produced by a set of lenses. The resolving power offers a quantifiable means of measuring the ability of a lens system to resolve detail and is defined as the smallest distance between two points that can still be distinguished as two separate entities. Although the calculation of resolution is quite complex and depends on a number of factors, the real world (as opposed to theoretical) limits of resolution can be calculated using the following equation: R = 0.61λ /NA where R refers to the resolving power of the optical system, λ is the wavelength of the light used, and NA is the numerical aperture of the objective being used. The numerical aperture describes the ability of a lens to gather light and resolve fine specimen detail at a fixed object distance. Numerical aperture is dependent on several aspects of the lens and is quite complex to calculate; it usually ranges from 0.4 (low power) to 1.25 (oil immersion) on most microscopes. Fortunately, numerical aperture, like magnification, is inscribed on the barrel of each objective lens (Figure 2.3). Resolution is also dependent on the wavelength of light used for examining a specimen, and as the equation presented earlier reveals, when the wavelength of light decreases, the resolving power gets smaller. Most microscopes use filters to produce blue or green light because these wavelengths are among the shortest seen clearly by humans. Plugging numbers into the equation shows that, using a wavelength of 500 nm and a lens with a numerical aperture of 1.25, the maximum resolution obtainable will be about 244 nm (0.244 µm). This means that two points on a specimen that are less than 0.244 µm apart are not resolvable and will be seen as a single entity. Ensuring that enough light passes through the sample to produce an adequate image is more complicated than one would initially think. Four factors go a long way toward guaranteeing that your images have maximum resolution and adequate contrast.

∙∙ Blue Light  The shorter wavelengths produced when a blue filter is placed over the light source will increase the resolution of the lens. ∙∙ Condenser Position  The condenser should be raised to its uppermost position. This maximizes the amount of light entering the objective lens and minimizes the amount lost to refraction. ∙∙ Diaphragm Position  The diaphragm should be closed just enough to provide an acceptable image. Although closing the diaphragm increases the contrast, it also decreases the numerical aperture. The best results are usually obtained by beginning an examination on low power with the diaphragm almost completely closed. As higher power objectives are inserted into the light path, open the diaphragm to provide more light. By the time the oil immersion lens is in place, the diaphragm should be almost completely open to maximize resolution. In general, the voltage control should only be adjusted when the image is too dark with the diaphragm completely open. ∙∙ Immersion Oil  This clear mineral oil has the same refractive index as glass and, when used between the slide and oil immersion lens, prevents the loss of light rays due to refraction (Figure 2.4). Recall that immersion oil is only to be used with the oil immersion lens. If oil gets on other lenses, the lens should be cleaned with lens paper dampened with alcohol. Be sure to clean oil from all the lenses when you have finished your observations.

OTHER LIGHT MICROSCOPES Brightfield microscopes are good general-purpose instruments and quite often the only microscope found in many laboratories. One disadvantage of this type of microscope is that images are generally poor if the specimen is lacking contrast. Although stains may be used to increase the contrast of microbial specimens, this almost always results in the death of the cell.



Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

(a)

(b)

Figure 2.5  Two views of a eukaryotic cell. This alga (Volvox) was examined while still alive using (a) brightfield (400x) and (b) darkfield (400x) microscopy. Notice the differences in the microscopic field and degree of detail afforded by each type of microscope. ©Stephen Durr Modification of the basic structure of a brightfield microscope has resulted in other types of light microscopes with unique properties, including the ability to produce good images from specimens with little contrast. A darkfield microscope blocks most of the light passing through the condenser so that only those light rays reflecting off of an object in the microscopic field are used to form the image. The result is a brightly illuminated object being viewed against a darkened field, the opposite of what is seen with a brightfield microscope (Figure 2.5). Phase-contrast microscopes employ a complex optical system in which retardation of the phase of light waves by cellular structures is converted to differences in the intensity of light, resulting in a high-contrast, detailed image that is especially useful when examining living (unstained) specimens.

MICROSCOPIC MEASUREMENTS In addition to a clear image of a specimen, it is sometimes desirable to have an accurate size measurement as well, and microscopes

13

can be easily outfitted with ocular and stage micrometers for this purpose. An ocular micrometer is simply a circular glass disc that has a series of regularly spaced markings etched onto its upper surface. The micrometer is installed within one of the oculars anytime a measurement is required, although in practice most laboratories find it easier to have an extra ocular with the micrometer permanently attached. When measurements are necessary, the ocular containing the micrometer is simply exchanged with one of the oculars in the microscope. The distance between the markings on an ocular micrometer have no meaning until the ocular is paired with an objective lens and calibrated using a stage micrometer. The stage micrometer resembles a slide except that it has markings etched upon it that are exactly 0.1 mm (100 μm) and 0.01 mm (10 μm) apart (Figure 2.6). To calibrate the ocular micrometer for use with a specific objective, the scales on the two micrometers must be superimposed on one another and the number of ocular graduations per stage graduations (10 or 100 μm) is then determined. If, when using the high power lens, for example, seven ocular divisions align with one stage division, then each ocular division equals 10  μm/7 ocular divisions or 1.43 μm/ocular division. This process is summarized in Figure  2.7. Once the ocular micrometer is calibrated for use with a specific objective lens, the stage micrometer is removed and replaced with a slide containing the organism to be measured. By counting the number of ocular graduations and multiplying by the distance between the graduations, the size of an unknown specimen is easily calculated. Because of the thickness of the stage micrometer, it is generally impossible to directly calibrate an ocular micrometer for use with the oil immersion lens. In this case, calibration can be accomplished mathematically. Using the example from the previous paragraph, one ocular division equals 1.43 μm when viewed using the high power (40x) objective. When using the oil immersion (100x) objective, an object should appear 2.5 times larger than when viewed under high power (100/40 = 2.5), but the inscribed marks on the ocular haven’t changed at all. Therefore, the distance between two marks on the ocular micrometer now spans a distance 2.5 times as great as the same marks did under the high power lens. To calibrate the ocular micrometer for use with the oil immersion lens, all that is needed is to divide the size of each ocular division (1.43 μm in the previous example) by the difference in the magnification of each lens (100/40 or 2.5): 1.43 μm per ocular unit (high power)/2.5 = 0.57 μm (oil)

Figure 2.6  A stage micrometer has markings etched onto its surface that are a precise distance from one another. The label on the micrometer, in this case 0.01 mm, indicates that the distance between two adjacent lines is 0.01 mm (10 μm). ©Aaron Roeth Photography

14

Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

X

Y

Y

(a) View showing the alignment of stage micrometer graduations (X) with ocular micrometer graduations (Y). Since one space of X (0.01 mm) is occupied by seven spaces of Y, one space of Y .01 = .0014 mm, or 1.4 micrometers. 7

(b) View showing appearance of ocular micrometer graduations. Spacing is arbitrary.

X

(d) On the basis of the calibration calculations in view (a) above, the budding yeast cell seen here is (15 units x 14 µM/unit) 21 µM in length.

(c) Appearance of stage micrometer graduations. Lines are exactly 0.01 mm (10 micrometers) apart.

Figure 2.7  Procedure for the use of stage and ocular micrometers to measure microscopic specimens.

PRE-LAB QUESTIONS

Remember, Understand, Apply 1. When using a brightfield microscope, the amount of light entering the objective lens is primarily regulated by the a. ocular lenses. b. diaphragm. c. mechanical stage. d. condenser. 2. When transporting a microscope, it should be carried by the a. base and condenser. b. arm and diaphragm.

c. base and arm. d. ocular and objective lenses. 3. The most common cause of an unacceptable microscopic image is a. an improperly adjusted condenser. b. insufficient light. c. an objective lens that is not matched to the ocular lens. d. dirty lenses. 4. Objective lenses should always be coated in immersion oil to keep them protected against dust and dirt. a. true b. false



Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

5. What can be safely used to clean microscope lenses? a. cotton swabs b. a clean, cotton cloth c. a paper towel d. a Kimwipe or other laboratory tissue 6. The total magnification of a microscopic image can be calculated by using which of the following formula? a. 0.61λ/NA b. magnification of the ocular lens/by numerical aperture c. resolution of the objective lens/magnification of the objective lens d. magnification of the ocular × magnification of the objective 7. The resolution of a microscopic image refers to a. how detailed the image appears. b. how well-focused the image appears. c. how large the image appears. d. how much contrast the image possesses. 8. Immersion oil improves the quality of a microscopic image by removing which of the following from the light path? a. air b. water c. oil d. dirt 9. A microscope which produces a light image against a dark background is most likely a a. brightfield microscope. b. phase-contrast microscope. c. darkfield microscope. d. electron microscope. 10. Complete each of the tables below.

Total magnification

Ocular magnification

750x

10x

10x

10x

45x

10x

100x

10x

225x

Resolution

Objective magnification

45x

Wavelength

Numerical aperture

500 μm

 0.4

700 μm

 0.4

500 μm

1.25

15

MATERIALS Each student should have: A microscope Prepared slides of: Bacteria Vorticella or Spirogyra Paramecium Access to stage and ocular micrometers

PROCEDURE Part I: Observations 1. Carefully carry your microscope to your work area and place it gently on the benchtop. 2. Plug in the microscope and turn on the light source, keeping the voltage control (brightness) at a minimum. Be sure the condenser is raised to its maximum and the diaphragm is almost completely closed. Increase the voltage to the lamp until the illumination is at a comfortable level. 3. Place a slide on the stage, holding it in place with the stage clips. Be sure that the specimen is on the upper surface of the slide. If the specimen is on the lower surface of the slide, you will be able to focus when using low power and high dry, but not when using the oil immersion lens. 4. Center the slide over the light beam emanating from the condenser. Move the lowest power objective (usually 10x) into position, listening for an audible click as it slips into place. 5. Bring the image into focus using first the coarse focusing knob and then the fine focusing knob to obtain the sharpest possible image. If you are using a binocular microscope, use the diopter adjustment to compensate for differences in visual acuity between your eyes as follows: ∙∙ Close the eye with the adjustable ocular, and focus for your open eye using the coarse and fine adjustment knobs. ∙∙ Using only the eye with the adjustable ocular, turn the diopter adjustment until the image is in sharp focus. 6. Adjust the interpupillary distance to match the distance between your eyes by looking though both oculars and slowly adjusting the distance between them until a single image is seen. If you will be measuring specimens in today’s lab, follow the instructions in Part II to calibrate the ocular micrometer before proceeding. Otherwise, continue with the steps that follow. 7. Place a prepared slide on the stage of your microscope. Use the mechanical stage to hold it in place. 8. Scan the slide using the low power objective, selecting a potentially interesting area for further examination in greater detail. 9. Move the slide so that the area you wish to examine is centered in the microscopic field. Your microscope is parcentric, meaning that once a specimen is located in the center of the field, it will remain centered when changing objectives.

16

Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

10. Swing the next highest objective into place, making sure that it clicks into position. Most microscopes are parfocal, meaning that if a specimen is focused with one objective, the image should remain sharp as the objectives are changed. You will have to make small adjustments to both the fine focus and diaphragm for each new objective. 11. When viewing bacteria, you will need to use the oil immersion lens. After viewing the specimen with the high dry lens and obtaining a clear image, swing the lens out of the way and add a single drop of immersion oil to the slide directly above the condenser. Rotate the oil immersion (100x) lens into place, making sure that the end of the lens is submerged in the oil. Open the diaphragm almost all the way to get the highest quality image.

If you “lose” your sample after changing lenses, it is best to go back to low power and begin again. If you’ve already added oil to the slide, go straight from low power to oil immersion so you won’t get oil on the high dry lens. 12. For each of your slides, record drawings of the specimens you observed in the space provided. Label each drawing with the name of the sample, total magnification, and size if measurements are being made. 13. When you are done for the period, clean any oil off the lenses of your microscope as well as off the stage, focusing knobs, prepared slides, etc. 14. Rotate the low power objective into position, center the mechanical stage, lower the lamp’s voltage to a minimum, and switch it off.

15. Wrap or gather the electrical cord according to the rules of your lab and return your microscope to its storage area. Part II: Measurements 1. Following the instructions for your particular laboratory, insert an ocular containing a micrometer into your microscope. 2. Move the lowest power objective into place, making sure it clicks into position. 3. Place a stage micrometer onto the stage and hold it in place with the stage clips. Move the stage micrometer, and rotate the ocular micrometer until the two are superimposed on one another with the left sides of each aligned. 4. Record the number of ocular gradations per stage gradations in the accompanying table. 5. Repeat the process with the high power lens in place. 6. Repeat the process with the oil immersion lens, or calibrate the ocular micrometer for use with the oil immersion lens mathematically.

Lens

Ocular units (OU)

Distance (from stage micrometer)

Distance/OU

Scanning Low power High dry Oil immersion

7. After completing calibration of the ocular micrometer, return to the procedure above.

RESULTS Record your results. For each specimen, provide as accurate a drawing as possible. Include the total magnification used to observe the specimen (i.e., 100x, 450x, or 1000x) and indicate the size of each specimen if measurements were taken. Specimen ____________    (______X)

Specimen ____________    (______X)

Specimen ____________    (______X)

Specimen ____________    (______X)



Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

REVIEW QUESTIONS

Analyze, Evaluate, Create 1. What is meant by the terms parcentric and parfocal?

2. In order to make specimens easier to see, why don’t microscope makers use 100x ocular lenses?

3. What is the total magnification of a microscope with a 45x objective lens and a 15x ocular?

4. Describe how the condenser and diaphragm should be adjusted for optimum viewing of a specimen using the oil immersion lens.

5. Describe how to properly clean the objective lenses of a microscope.

6. For a microscope on which 17 ocular units align with 100 μm on the stage micrometer when the 4x objective is used, how many micrometers are there per ocular unit when using the (a) 4x objective, (b) 10x objective, (c) 45x objective, and (d) 97x objective?

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Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

CASE STUDY Prior to the middle of the 19th century, there was no science of microbiology, because organisms so small as to appear invisible to the naked eye were not thought to exist. Of course, no microbiology meant no microbiologists. Persons who found interest in the natural world were often amateurs, little different from the backyard astronomer who discovers a new comet or the birdwatcher who discovers a new species by accident—talented people interested in the world around them. One of these amateurs lived in the Netherlands in the 1600s. Anton van Leeuwenhoek, a fabric salesman by trade, had an interest in glass production and became an expert

(a)

(b)

at making small lenses, creating by far the most powerful microscopes of the time. Like the backyard astronomer, he also had an interest in the world around him, and he used his lenses to examine water pulled from lakes, mold on bread, even the plaque between his teeth. He recorded his observations and sent them to the Royal Society of London for the Improvement of Natural Knowledge (an organization which still exists, see www.royalsociety.org). Over the course of many letters, Leeuwenhoek described “. . .green streaks, spirally wound serpentwise.  .  .”, “.  .  .little animals fashioned like a bell. . .”, and from his own teeth “. . .many very little living animalcules, very prettily a-moving.”

(c)

Figure 2.8  (a) Spirogyra, an alga; (b) Vorticella, a protozoan; and (c) dental plaque, consisting of various species of bacteria. (a) ©Stephen Durr;

(b) ©McGraw-Hill Education/Richard Gross, photographer; (c) ©Science Photo Library-STEVE GSCHMEISSNER./Getty Images

Lens Specimen holder

Focus screw

Handle

A brass replica of one of Leeuwenhoek’s original microscopes. ©Kathy Park Talaro

While Leeuwenhoek was initially credited with being an adept observer of the microscopic world around him, describing the structure of wood, a honeybee, or body lice, his observations of living microorganisms caused his credibility to be called into question as such things were “known” not to exist. After an extensive investigation in the 1670s, it was the Royal Society who rethought their own theories of life, based on Leeuwenhoek’s observations. Anton van Leeuwenhoek is considered one of the very first microbiologists, using the microscopes he built himself to examine a previously invisible world. Compare what you’ve seen today with what he saw, and described, 300 years ago. From Leeuwenhoek’s descriptions as well as detailed examination of his samples made with modern instruments, it is thought that the organisms he described as “green streaks, spirally wound .  .  .” were the alga Spirogyra, those “fashioned like a bell .  .  .” were the alga Vorticella, and the inhabitants of plaque were various species of bacteria (Figure  2.8). Leeuwenhoek’s microscopic examination of everyday items and his descriptions of the microorganisms he found occurred over 150 years before the so-called Golden Age of Microbiology. Algae similar to those he saw are still tracked today because of their potential to cause a



Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

variety of neurological illnesses and alter the biology of lakes and oceans. His description of the myriad bacteria found between his own teeth is probably the first ever description of a biofilm, a type of bacterial community essential to the formation of dental caries (cavities) and a number of other

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diseases. Leeuwenhoek’s correspondence with the Royal Society of London spanned more than 50 years. Prior to his death in 1723, he was credited with being the first to observe protozoa, bacteria, spermatozoa, and the banded pattern of muscle fibers.

CASE STUDY ANALYSIS 1. Leeuwenhoek was credited with being among the first to describe many biological entities. Nearly 300 years later, the University of Utah designed a web application that compares the sizes of different biological objects (http://learn.genetics.utah.edu/content/cells /scale/). Use the website to draw, to scale, an amoeba, paramecium, sperm cell, red blood cell, yeast cell, and E. coli cell. Be sure to label each cell with its approximate size.

REFERENCE University of California Museum of Paleontology. Antony Leeuwenhoek (1632–1723). www.ucmp.berkeley.edu/history/leeuwenhoek.html.

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Case Study Exercise 2  Microscopy and Measurement of Microscopic Specimens

NOTES

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C A S E S T U DY E X E R C I S E

A Survey of Protists STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Prepare a wet mount of a liquid sample. 2. Differentiate unicellular eukaryotes (protists) based on physical appearance and physiological traits.

INTRODUCTION Rather than being a well-defined taxonomic group with a clear evolutionary history, the protists are a diverse collection of organisms with evolutionary roots spread across the Domain Eukarya. Historically, the Kingdom Protista included most single-celled eukaryotes and was divided into two large subkingdoms. Those organisms that obtained nutrition through photosynthesis—were in essence plant-like—were assigned to the subkingdom Algae, while organisms that ingested organic matter—acting like animals— were assigned to the subkingdom Protozoa (Table 3.1). While this system of classification was simple and deceivingly easy to use (seaweed looks like a plant, right?), it was also incorrect. As

TABLE 3.1

molecular and genetic analysis began to play a larger role in the identification and taxonomy of organisms, this classification of protists fell apart. As currently classified, the protists include any single-celled organism that is not a plant, animal, or fungus. Beyond that, any two randomly selected protists may seem to have little in common. Rather than spending a great deal of time memorizing a traditional Kingdom/Phylum/Class taxonomic organization, we will use a more informal system in which protists are gathered into supergroups, groups, and subgroups. This is not an uncommon strategy because the characteristics of individual members are more important (and far more interesting) than the relationships between the hundreds of thousands of protist species.

Supergroup Excavata Most members of this supergroup are asymmetrical and have a feeding groove that appears to be excavated into one side of the organism. Two of the three groups within the excavata—the diplomonads and the parabasalids—possess highly reduced (i.e., highly simplified and poorly functioning) mitochondria and generate most of their energy anaerobically. The relationship between the members of the excavata can be seen in Figure 3.1.

Traditional Classification of Unicellular Eukaryotes Single-celled Eukaryotes

Subkingdom Protozoa2 (heterotrophic)

Subkingdom Algae1 (photosynthetic)

Excavata

Archaeplastida

Chromalveolata

Euglenoids

Green algae Red algae

Dinoflagellates Synura Yellow-green algae Golden-brown algae Brown algae Diatoms

Flagellates

Amoebae

Ciliates

Apicomplexa

1. Algae are further classified by the type of photosynthetic pigments they contain, the type of cell wall or covering they possess, and the type of molecule they use to store energy. 2. Protozoans are further classified by the type of motility they display. Flagellate (via flagella), Amoeba (via pseudopods), Ciliates (via cilia). Apicomplexans are typically nonmotile.

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Case Study Exercise 3  A Survey of Protists

Supergroup Excavata

Diplomonads

Parabasalids

Euglenozoans

Giardia

Trichomonas

Euglena Trypanosoma

Figure 3.1  Organization of the supergroup Excavata. The supergroup is shaded dark blue, groups are shaded medium blue, and common representative genera are unshaded.

Giardia duodenalis (also known as G. lamblia or G. intestinalis) is a diplomonad found throughout the world (Figure 3.2). Giardia cells may encase themselves in tough outer coverings called cysts, which allow them to survive harsh environmental conditions, including the low pH of the stomach. When Giardia cysts are ingested, through water contaminated with feces from infected people or animals, or as a result of poor hygiene, the cysts travel through the stomach to the small intestine, where excystation

Figure 3.2  Electron micrograph of Giardia duodenalis in the process of cell division. The nearly doubled pear-shaped organism currently resembles a heart. This species has four pairs of flagella for attaching to the intestinal epithelium (2500x). Source: CDC/Dr. Stan Erlandsen

produces two trophozoites (the active, proliferative form of the organism) per cyst. Infections may be asymptomatic, but most are marked by severe diarrhea lasting as long as six weeks. Infected persons periodically release large numbers of cysts into their feces, spreading the organism. Infection can be diagnosed by observing the stool for cysts or through the use of sensitive serological assays. Members of the second subgroup in the excavata, the parabasalids, also have reduced mitochondria. A common example is Trichomonas vaginalis (Figure 3.3), the cause of trichomoniasis in humans, a disease marked by inflammation of the genitourinary tract. An important difference between these two parasites is that T. vaginalis does not form cysts, leaving the more delicate trophozoite as the infective form. Because the trophozoite is far less hardy than a cyst, T. vaginalis must be passed directly from person to person, causing trichomoniasis to be classified as a sexually transmitted disease. The third group of the excavata is the euglenozoans, which are characterized by a unique crystalline structure within their flagella. This group is divided into two subgroups, the kinetoplastids and the euglenids. Named for a mass of DNA within their single mitochondria called a kinetoplast, the kinetoplastid group includes free-living heterotrophs as well as parasites. The most medically important species are in the genera Trypanosoma and are spread by the bite of bloodsucking insects. Trypanosoma brucei (Figure 3.4) causes sleeping sickness, a neurological disease that is fatal if left

Figure 3.3  Trichomonas vaginalis. With no cyst form, the parasite does not survive well outside the host, and it is transmitted almost exclusively through sexual intercourse (800x). Source: CDC/Dr. Mae Melvin



Case Study Exercise 3  A Survey of Protists

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Figure 3.4  Trypanosoma brucei parasites within a blood smear (the small, pink, round objects are red blood cells). T. brucei is the cause of sleeping sickness and is spread by the Tsetse fly, a bloodsucking insect (1000x). Source: CDC/Dr. Mae Melvin

Figure 3.5  Euglena sp. The red eyespot, flagellum, and translucent grains of paramylon are all visible in this photo (400x). ©Lebendkulturen. de/Shutterstock

untreated, while Trypanosoma cruzi causes Chagas’ disease and can lead to cardiac failure. Most euglenids are mixotrophs, meaning they may act photosynthetically (deriving energy from sunlight) or heterotrophically (deriving energy from the consumption of organisms). Euglena possesses a red eyespot at one end of the cell that detects light, along with one or two long flagella for motion. Together the eyespot and the flagella allow the organism to move to areas with the best light intensity for photosynthesis. Euglena stores energy from sunlight as paramylon, a lipopolysaccharide. When light is absent, Euglena behaves as a heterotroph, absorbing organic nutrients from the environment or preying on smaller organisms (Figure 3.5).

Supergroup SAR This large group contains organism that often don’t look or act alike, yet have been shown to be closely related based on sequencing of their DNA. With no single name accepted by the scientific community, the supergroup SAR is an acronym for the three

primary groups found within: the stramenopiles, the alveolates, and the rhizarians. The relationship between the members of the SAR supergroup is illustrated in Figure 3.6.

Stramenopiles Stramenopiles are characterized by a single unusual flagellum, which is covered with short, hair-like projections. Most species also have a second flagellum that is both shorter and smoother. The stramenopiles are divided into three subgroups: the diatoms, golden algae, and brown algae. Stramenopiles typically share a certain degree of greenish-brown color due to the green chlorophyll, brown fucoxanthin, orange beta-carotene, and yellow xanthophyll photosynthetic pigments they possess. Diatoms are a large group (100,000 species) of unicellular algae. These organisms have outer shells called frustules composed of silicon dioxide embedded in an organic matrix. The frustule consists of two halves, with the larger half, or epitheca, fitting over the smaller half, or hypotheca, like the top and bottom of a Petri dish. Diatoms are some of the most numerous organisms in

Supergroup SAR

Stramenopiles

Diatoms

Golden algae

Alveolates

Brown algae

Ciliophora

Dinoflagellates

Rhizarians

Apicomplexans

Foraminiferans

Radiolarians

Figure 3.6  Organization of the supergroup SAR. The supergroup is shaded dark blue, groups are shaded medium blue, and subgroups are shaded light blue.

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Case Study Exercise 3  A Survey of Protists

(a) Diatom. Scanning electron micrograph (1600x).

(d) Diatom (10,000x). Colored scanning electron micrograph of a cell wall of a diatom.

(b) Didymosphenia geminata (800x). Collected from the Rio Espolon in Chile during an algal bloom and processed to display the silica cell wall, Didymosphenia is considered an invasive species.

(e) Diatom (800x). Colored scanning electron micrograph of a cell wall of a diatom.

(c) Licmorpha sp. (500x). A freshwater diatom.

(f) Odontella mobiliensis (400x). A marine diatom isolated from Long Island Sound.

Figure 3.7  Diatom gallery. (a, c-e, h, l) ©Science Photo Library/Alamy Stock Photo; (b) Source: Sarah Spaulding/USGS; (f, i, k) Source: Dr. Yaqin ‘Judy’ Li, Milford Laboratory/ NEFSC/NMFS/NOAA; (g) ©Oxford Scientific/Getty Images; ( j) ©Steve Gschmeissner/Science Source

the sea and serve as a primary food source for filter feeders like clams, oysters, and mussels, along with other marine organisms. They store energy from photosynthesis as chrysolaminarin, a polysaccharide, or oil, and many scientists believe that our worldwide petroleum reserves are the result of the photosynthetic actions of diatoms over millions of years. After death, the frustules of dead diatoms sink to the bottom of lakes and oceans where they form deposits of material called diatomaceous earth, used industrially in polishes, abrasives, and water filters. These dead diatoms also take with them to the ocean floor all of the carbon in their bodies, which remains trapped for decades, perhaps centuries. If the diatoms had instead been ingested near the surface, the carbon they contained would be released almost immediately as carbon dioxide (CO2). The worldwide diatom population is so large that the amount of CO2 “pumped” to the ocean floor may measurably reduce global carbon dioxide levels. The unique structure of diatoms can be seen in Figure 3.7. Golden algae is a large subgroup containing over 6000 species. A preponderance of xanthophylls and fucoxanthin give most species in this group a golden-brown appearance. Some

species of golden algae are mixotrophic and can absorb organic compounds and phagocytize living cells. Poor environmental conditions stimulate the formation of protective cysts, which can allow survival for decades. Common members include synura (Figure 3.8). The brown algae subgroup consists almost entirely of multicellular marine organisms, and most seaweeds are members of this group. A greater amount of fucoxanthin and carotenoids lends these organisms a decidedly brownish hue. Algin, a polysaccharide extracted from the cell wall of brown algae, is used as a thickening agent in foods and cosmetics and to reduce ice crystal formation in ice cream. Food storage is in the form of laminarin, a polysaccharide, and mannitol, a sugar alcohol. Several examples of brown algae may be seen in Figure 3.9.

Alveolates Named for the presence of membrane-enclosed sacs called alveoli which lie below the cell membrane, the alveolates include three subgroups: the dinoflagellates, the apicomplexans, and the ciliates.



(g) Diatom frustules (400x).

( j) Coscinodiscus sp. (600x). Colored scanning electron micrograph.

Figure 3.7  Diatom gallery. (continued)

Case Study Exercise 3  A Survey of Protists

(h) Diatom frustules (800x). Colorized scanning electron micrograph of a diatom frustule, emphasizing the silicon dioxide cell wall.

(k) Gyrosigma sp. (wider species on bottom) and Pseudo-nitzschia sp. (thin specimen on top) (400x). Several species of Pseudo-nitzschia produce the neurotoxin domoic acid, responsible for amnesic shellfish poisoning in humans.

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(i) Eucampia zoodiacus (400x). A marine diatom isolated from Long Island Sound.

(l) Cylotella sp. (3,500x). A diatom found in both fresh and marine waters. Cylotellawas first seen in Lake Michigan in 1964 but has since spread to all of the Great Lakes.

Dinoflagellates are a group of unicellular organisms commonly found in marine waters. Most dinoflagellates are covered by two protective plates and possess two flagella, each of which rests within a groove encircling the organism (Figure 3.10). When moving, the longitudinal flagellum extends behind the cell while the transverse flagellum beats within its groove, spinning the cell as it is propelled forward. It is from this unique motion that the dinoflagellates received their name (from the Greek dini, “to whirl”). Roughly half of all dinoflagellates are heterotrophic, with the rest being mixotrophic or photosynthetic. Photosynthetic dinoflagellates typically possess chlorophylls a and c, as well as beta-carotenes and xanthophylls, which lend an orange color to

Figure 3.8  Synura owes its golden color to a combination of fucoxanthin and xanthophyll (400x). Not visible are the flagella that allow Synura to position itself where the light intensity is greatest, thereby maximizing photosynthetic activity. ©McGraw-Hill Education/Stephen Durr, photographer

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Case Study Exercise 3  A Survey of Protists

(a)

(b)

Figure 3.9  Brown algae. (a) Fucus, a small seaweed (up to 2 m) found in intertidal zones throughout the world. (b) Macrocystis pyrifera (giant kelp) may grow to over 200 m in length. The air bladders visible at the base of each blade provide buoyancy, lifting the kelp toward the surface of the water and allowing it to gather more light for photosynthesis. (c) Alaria marginata grows to 4 m in length and is found in the intertidal zone. Because it only grows in shallow water, air bladders, like those seen in Macrocystis, are not needed. (c)

(a) ©McGraw-Hill Education/Richard Gross, photographer; (b) ©Claire Fackler, CINMS, NOAA; (c) Source: Mandy Lindeberg, NOAA/NMFS/AKFSC

water during population blooms known as red tides (Figure 3.11), sometimes poisoning fish and other marine organisms that feed on the algae. Depending on the species, foods are stored as starches or oils. The apicomplexans consists of organisms that share a unique structure called an apical complex used for penetrating host cells.

Nearly all members of the group are parasitic and have complex life cycles which include multiple hosts. A prime example of this group is Plasmodium, the causative agent of malaria, a disease which kills upwards of 200,000 people each year. Five species of Plasmodium cause malaria in humans: P. falciparum, P. malariae, P. ovale, P. vivax, and P. knowlesi. The life cycle is split between



(a)

Case Study Exercise 3  A Survey of Protists

(b)

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(c)

Figure 3.10  (a) and (b) Ceratium (400x), common members of the alveolata (dinoflagellates). Notice the position of the flagellum within a groove encircling the organism. (c) Peridinium, another common dinoflagellate. (a) Source: Steve Morton, Marine Biotoxins Program, NOAA

structure. Ciliates are unique in possessing two types of nuclei: micronuclei and macronuclei. Genes in the macronucleus control what are colloquially known as “housekeeping” functions, such as feeding and waste removal, while the micronuclei are exchanged between ciliates in a process called conjugation, resulting in genetic variability. The vast majority of ciliates are free living and harmless and generally exhibit the most complex structures and behaviors of all protists (Figure 3.15). The most commonly encountered parasite is Balantidium coli, which can cause bloody and mucus-filled diarrhea, although most infections are thought to be asymptomatic.

Rhizarians Figure 3.11  Red tide along the coast of Puget Sound. The combination of warm water, fertilizer runoff, and long periods of daylight leads to algal blooms. Toxins produced by the algae are concentrated in the bodies of filter-feeding organisms like clams and mussels, eventually working their way up the food chain to large fish, birds, and humans. ©Don Paulson/SuperStock/Corbis

human tissues (red blood cells and liver) and an insect vector, the female Anopheles mosquito (Figure 3.12). Diagnosis, as well as determination of the infectious species, can be accomplished by examining red blood cells for the presence of the parasite (Figures 3.13 and 3.14). Ciliates are named for the numerous small, hair-like projections covering the cell, providing the cell with motility and, in some cases, a means of directing food toward a specialized feeding

Rhizarians are the “R” in SAR. Most members of this subgroup are amoebas with thread-like pseudopods, cellular extensions of the cell used in motility and feeding. As a pseudopod extends from the cell, cytoplasm streams into the growing appendage, moving the cell in that direction. Alternatively, food particles that have become attached to the tip of a pseudopod can be carried into the cell as the pseudopod retracts. Rhizarians are divided into two large subgroups: radiolarians and foraminiferans (forams). The skeletons associated with these two groups can be exquisitely beautiful and are seen in Figure 3.16.

Radiolarians Members of this group of rhizarians are marked by delicate, symmetrical, internal skeletons composed of silica. Hair-like pseudopods radiate from the central part of the cell and are used for motility or to procure food.

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Case Study Exercise 3  A Survey of Protists i = Infective Stage d = Diagnostic Stage

Human Liver Stages Liver cell

2 Infected liver cell

Mosquito Stages 12 Ruptured oocyst 11 Oocyst i

1 i Mosquito takes a blood meal (injects sporozoites)

Release of sporozoites

4 Ruptured schizont

C Sporogonic Cycle

10 Ookinete

Macrogametocyte

3 Schizont

A Exo-erythrocytic cycle

5

Human Blood Stages

d Immature trophozoite (ring stage)

8 Mosquito takes a blood meal (ingests gametocytes) B Erythrocytic Cycle

9 Microgamete entering macrogamete

d Mature trophozoite

P. falciparum O +

Exflagellated microgametocyte O +

P. vivax P. ovale P. malarian

O

6 Ruptured schizont

7 Gametocytes d O

Schizont d 7 Gametocytes

Figure 3.12  Life cycle of malaria. Malaria is a complex disease that is responsible for about 600,000 deaths each year worldwide. The disease may be caused by at least five species of Plasmodium (P. falciparum, P. vivax, P. ovale, P. malariae, and P. knowlesi), involves two hosts, and has a complex life cycle composed of three distinct components. An understanding of the life cycle is necessary when diagnosing the disease. During a blood meal, a malaria-infected female Anopheles mosquito inoculates sporozoites into the human host 1 . Sporozoites infect liver cells 2 and mature into schizonts 3 , which rupture and release merozoites 4 . After this initial replication in the liver (exoerythrocytic cycle A  ), the parasites undergo asexual multiplication in the erythrocytes (erythrocytic cycle B  ). Merozoites infect red blood cells 5 . The ring-stage trophozoites mature into schizonts, which rupture, releasing merozoites 6 . Some parasites differentiate into sexual erythrocytic stages, producing male and female gametocytes. The gametocytes, male (microgametocytes) and female (macrogametocytes), are ingested by an Anopheles mosquito during a blood meal 8 . The parasites’ multiplication in the mosquito is known as the sporogonic cycle C . While in the mosquito’s stomach, the microgametes penetrate the macrogametes, generating zygotes 9 . The zygotes in turn become motile and elongated ookinetes 10 , which invade the midgut wall of the mosquito where they develop into oocysts 11 . The oocysts grow, rupture, and release sporozoites 12 , which make their way to the mosquito’s salivary glands. Inoculation of the sporozoites into a new human host perpetuates the malaria life cycle 1 . (a-b) Source: CDC/Steven Glenn, Laboratory Training & Consultation Division



Case Study Exercise 3  A Survey of Protists

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(a) (b)

(c)

(d)

Figure 3.13  Stages of the Plasmodium falciparum erythrocytic cycle. (a) Ring-form trophozoites (1150x). This stage represents the asexual, immature stage of P. falciparum. Note the central red blood cell, which contains three trophozoites. (b) Ring forms as well as crescent-shaped gametocytes are seen in this Giemsa-stained blood smear (1200x). (c) Development of gametocytes leads to the production of a schizont (1000x) filled with as many as 24 merozoites. (d) Continued development of gametocytes leads to the production of microgametocytes (male) and megagametocytes (female). Here the sausage-shaped macrogametocyte is visible (1150x). (a, c, d) Source: CDC/Steven Glenn, Laboratory Training & Consultation Division; (b) CDC/ Dr. Greene; Steven Glenn, Laboratory & Consultation Division

Figure 3.14  Comparison of trophozoite stage of three Plasmodium species (1150x). As the trophozoite matures, its morphology changes, transitioning from ring form to mature trophozoite. The morphology of the mature trophozoite stage allows species identification of Plasmodium. (a) Plasmodium vivax. (b) Plasmodium malariae. (c) Plasmodium ovale. Source: (a)

(b)

(c)

CDC/Steven Glenn, Laboratory Training & Consultation Division

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Case Study Exercise 3  A Survey of Protists

(a) Paramecium caudatum (400x). The cilia visible on the periphery of the organism allow for motility and also direct food toward the oral groove, visible in the bottom center of the cell.

(b) Paramecium caudatum (430x) in the process of conjugation. During conjugation, two individual cells partially fuse and exchange micronuclei before separating, leading to increased genetic variation.

(d) Stentor coeruleus (50x). An exclusively freshwater protist which may grow to 2 mm in length, Stentor is one of the largest single-celled organisms yet identified.

(c) Vorticella sp. (430x). The edge of the vase-shaped cell body is ringed with cilia which beat to draw food closer to the organism. The stalk will alternately contract and extend to move the cell body (as seen in the inset), a behavior that is both defensive and increases food capture.

Figure 3.15  Ciliate gallery. (a) ©NNehring/Getty Images; (b) ©McGraw-Hill Education/Richard Gross, photographer; (c1) ©McGraw-Hill Education/Richard Gross, photographer; (c2) ©McGraw-Hill Education; (d) ©Oxford Scientific/Getty Images



(a)

Case Study Exercise 3  A Survey of Protists

(b)

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(c)

Figure 3.16  (top) Scanning electron micrographs of radiolarian skeletons (400x). (bottom) Scanning electron micrograph of foraminiferan tests (400x). (a-c) ©Science Photo Library/Alamy Stock Photo; (d)

(d) ©PASIEKA/SPL/Getty Images

Forams Forams are recognized, and in fact named for, the porous shells called tests, which surround them. Foram tests consist of a single piece of organic material hardened with calcium carbonate. Pseudopods within the test extend through pores in the shell and are used for motility and feeding.

Supergroup Archaeplastida

Supergroup Archaeplastida This large supergroup (Figure 3.17) contains all of the terrestrial plants along with their closest algal relatives, the red algae and green algae. The green plants are left for a future course in botany, while a discussion of algae follows.

Green Algae This large group containing over 7000 species is commonly seen in ponds, neglected swimming pools, and fish tanks. They are a diverse group with members found in fresh and salt water that

Red algae

Green algae

Rhodoglossum Gelidium

Chlorophyta Chlamydomonas Volvox Caulerpa

Figure 3.17  Organization of the supergroup Archaeplastida. The supergroup is shaded dark blue, groups are shaded medium blue, and common representative genera are unshaded.

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Case Study Exercise 3  A Survey of Protists

(b) (a)

Figure 3.18   Green algae may exhibit a (a) unicellular (Micrasteria rotata 400x), (b) colonial (Gonium sp., 400x), or (c) filamentous (Hormidium sp., 400x) structure. (a) ©Lebendkulturen.de/Shutterstock; (b) ©McGraw-Hill Education/Stephen Durr, photographer; (c) ©Stephen Durr

(c)

exist both on their own and in association with other organisms. Molecular studies have shown the members of this group to be closely related to land plants. Like plants, members of the green algae group use chlorophylls a and b for photosynthesis (providing them with their characteristic bright green color), store the energy they create in the form of starch, and have cell walls made of cellulose. Members of this group may be unicellular, colonial, or filamentous in nature (Figure 3.18). Additional examples of green algae can be seen in Figure 3.19.

Red Algae This group, numbering more than 6000 species, owes its red color to the photosynthetic pigment phycoerythrin, whose presence often masks the green chlorophyll which is also present. Several members of the red algae are economically useful; the genus Porphyra serves as a wrap for sushi, while the genus Gelidium is the source of agar for microbiological media. A variety of red alga may be seen in Figure 3.20.



Case Study Exercise 3  A Survey of Protists

33

(a) Micrasterias truncata (400x). All members of the genus Micrasterias display bilateral symmetry, with two mirror image semicells joined by a narrow band containing the nucleus of the organism. Each semicell contains a single large chloroplast.

(b) Spirogyra conjugating (430x). Chloroplasts of Spirogyra are wound into a spiral within the cell, giving it a unique appearance. The algae is common in warm water where it forms filamentous masses, often joining together with the two other genera of algae, Mougeotia and Zygnema, to form large floating mats. The conjugation tubes connecting the two filaments of Spirogyra permit the exchange of genetic material, allowing sexual reproduction to occur.

(c) Volvox sp. (200x). Note the small daughter colonies within each of the main colonies.

(d) Caulerpa taxifolia. Commonly used in salt-water aquaria because of its hardy nature, Caulerpa quickly dominates when released into coastal waters. For this reason many states, including California, have banned its use.

(e) Zygnema sp. (100x). Zygnema species are composed of unbranched filaments with two stellate (star-shaped) chloroplasts in each cell.

(f) A green algae bloom in an Iowa pond.

Figure 3.19  Chlorophyta (green algae) gallery. (a) ©Lebendkulturen.de/Shutterstock; (b) ©McGraw-Hill Education/Richard Gross, photographer; (c) ©micro_photo/ Getty Images; (d-e) ©Steven P. Lynch; (f) ©Aaron Roeth Photography

(a) Rhodoglossum affine. A red algae occasionally found in the lower intertidal zone along the Pacific coast of the United States.

(b) Gigartina canaliculata. The red pigments found in members of the Rhodophyta allow them to photosynthesize far below the surface of the ocean.

0.1 mm

(c) Chondrus crispus. The combined effects of blue phycocyanin and orange caratenoid pigments create unique colors in many members of the rhodophyta.

(d) Polysiphonia sp. Found throughout the world but growing only to about 30 cm, Polysiphonia displays a complex, triphasic, lifecycle.

(e) Corallina vancouveriensis. This coralline (coral forming) red algae secretes lime, contributing to the formation of limestone reefs.

(f) Carrageenan. Many members of the red algae produce carrageenan, a thickening agent used in a variety of foods and consumer products.

Figure 3.20  Red algae gallery. (a, b, d-f) ©Steven P. Lynch; (c) ©Dan Ippolito



Case Study Exercise 3  A Survey of Protists

35

Supergroup Unikonta This diverse supergroup is the home to animals, fungi, and a few protists (the only group of interest to us, for now). All of the protists are found in the group amoebazoa and possess lobe-shaped pseudopods as opposed to the thread-like pseudopods seen in rhizarians. Amoebozoans are further classified into three subgroups: tubulinids, entamoebas, and slime molds (Figure 3.21). The tubulinid subgroup contains amoebas characterized by lobe-shaped pseudopods. Found throughout the world in soil, freshwater, and marine environments, these organisms are heterotrophs that typically prey on bacteria and protists for nutrition. Two views of common amoebas can be seen in Figure 3.22. Entamoebas differ from most other amoebas by being parasitic. Entamoeba histolytica is a serious pathogen of humans,

Supergroup Unikonta

Tubulinids

Amoebozoans

Animals

Entamoebas

Slime molds

Figure 3.23  Entamoeba histolytica trophozoite (400x). Source: CDC Fungi

Figure 3.21  Organization of the supergroup Unikonta. The supergroup is shaded dark blue, groups are shaded medium blue, and subgroups are shaded light blue.

(a)

responsible for nearly 100,000 deaths worldwide each year. The organism produces protective cysts which allow it to survive the hostile environment of the stomach prior to the release of trophozoites in the large intestine (Figure 3.23). Slime molds are large organisms (many centimeters in diameter) that exhibit complex life cycles and morphological changes.

(b)

Figure 3.22  (a) Amoeba proteus (160x). Several food vacuoles containing engulfed algae can be seen. (b) This scanning electron micrograph of the amoeba Korotnevella sp. (4000x) makes it easy to appreciate the three-dimensional morphology of the cell as well as the scales that cover the cell but that are impossible to see at lower magnification. (a) ©micro_photo/Getty Images; (b) ©Science Photo Library/Alamy Stock Photo

36

Case Study Exercise 3  A Survey of Protists

(a) Plasmodial slime mold. Although several cm across, this slime mold is undivided by a cellular membrane and contains multiple nuclei, making it a single cell.

(b) Hemitrichia serpula (pretzel slime mold).

(c) Fruiting bodies of Lycogala epidendrum.

(d) Fruiting bodies of Stemonitis fusca on a decaying log

Figure 3.24  Slime mold gallery. (a) ©McGraw-Hill Education/Richard Gross, photographer; (b) ©Mark Steinmetz; (c) ©Alexander62/iStock/Getty Images; (d) ©Ed Reschke/ Stockbyte/Getty Images

Slime molds are typically brightly colored and are commonly found in moist soil where they phagocytize dead organic material. A variety of slime molds can be seen in Figure 3.24.

Cyanobacteria Cyanobacteria are prokaryotic organisms that—because of their ability to photosynthesize along with their presence in aquatic environments—were for many years erroneously characterized as blue-green algae. These prokaryotic organisms are routinely found in lakes and ponds, and they vary greatly in shape and appearance (Figure 3.25). Cyanotoxins produced by cyanobacteria have the ability to cause serious illness, and in cities that draw drinking water from lakes, blooms of cyanobacteria have been the cause of water emergencies many times over the years.

Figure 3.25  Scanning electron micrograph of cyanobacteria. ©Science Photo Library - STEVE GSCHMEISSNER./Getty Images



Case Study Exercise 3  A Survey of Protists

PRE-LAB QUESTIONS

Remember, Understand, Apply 1. Which supergroup of protists contain members with a feeding groove on one side of the organism? a. excavata b. SAR c. archaeplastida d. unikonta 2. Characteristics important in the classification of members of the excavata include which two of the following? a. nucleus b. mitochondria c. pseudopod d. flagella 3. The protective covering that some protists can produce is a/n a. endospore. b. trophozoite. c. cyst. d. pseudopod. 4. Kelp (large seaweed) would most likely be found in which group within the SAR? a. stramenopiles b. apicomplexans c. alveolates d. rhizarians 5. A protist that may acquire energy both photosynthetically and heterotrophically is described as being a/n a. facultative phototroph. b. saprotroph. c. mixotroph. d. biphasic organism. 6. Members of the genus Plasmodium cause a. malaria. b. sleeping sickness. c. diarrheal disease. d. shellfish poisoning. 7. Diatoms have shells composed of a. cellulose. b. chitin. c. peptidoglycan. d. silicon dioxide. 8. The agar used to solidify plates in the microbiology lab is extracted from a. green algae. b. golden algae. c. brown algae. d. red algae.

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9. Human pathogens are found in which subgroup of the unikonta? a. tubulinid b. entamoeba c. slime molds d. poryphera 10. Long, threadlike pseudopods are found in members of which group? a. ciliates b. rhizarians c. alveolates d. stramenopiles 11. Match each of the photosynthetic pigments on the left with the colors on the right. a. chlorophyll ____ Red b. phycoerythrin ____ Orange c. carotenoids ____ Yellow d. xanthophyll ____ Green e. fucoxanthin ____ Brown 12. Define the following terms using the laboratory exercise or glossary: Flagellum

Cilia

Pseudopod

Photosynthetic pigment

MATERIALS Each student should have access to the following: Several bottles containing pond water samples Cultures of Amoeba Prepared slides of Trypanosoma sp. Prepared slides of Plasmodium sp. Microscope slides and cover slips Pasteur pipettes and forceps

38

Case Study Exercise 3  A Survey of Protists

PROCEDURE Make wet mount slides (one at a time) of each of the water samples available. A wet mount is a simple preparation method that allows the observation of live specimens. Prepare each wet mount as follows: 1. Use the pipette to extract a small amount of sample from the bottom of the sample jar. Very few organisms will be found in the middle of the sample. 2. If needed, use forceps to transfer a small amount of filamentous algae to a slide. 3. Place a cover slip gently on the sample. 4. Examine the slide under low power, looking for areas of interest. Adjust the diaphragm to reduce the light reaching

your slide, which will increase contrast and improve visibility. 5. When you encounter a specimen you would like to study in more detail, swing the high power (40x or 45x) objective into position. 6. The prepared slides of Trypanosoma and Plasmodium are blood smears taken from people or animals infected with the organism. Focus on the blood cells using the low and high power objectives on your microscope before attempting to find the microbial organism. Your instructor may want you to use the oil immersion lens for these prepared slides. If so, follow his or her instructions. 7. Use the pictures and descriptions found here, along with any supplementary books in your lab, to identify the organisms you encounter.

RESULTS Record your results. For each sample, provide a drawing that indicates the major features seen in the organism. Also indicate the total magnification used to observe the specimen (e.g., 100x and 450x). Based on the features you observe, identify each organism as best you can. Sample #1 (______X)

Sample #2 (______X)

Sample #3 (______X)

Sample #4 (______X)

Sample #5 (______X)

Sample #6 (______X)



Case Study Exercise 3  A Survey of Protists

REVIEW QUESTIONS Analyze, Evaluate, Create 1. What is the difference between a trophozoite and a cyst, and why is one generally more infective than the other?

2. A technician working for you in a microbial reference laboratory has examined a number of different samples and recorded his results. From his notes, identify the group or subgroup (be as specific as you can) represented by each description. a. A freshwater sample containing a colonial algae, bright green in color with cell walls made of cellulose.

b. A blood sample in which the red blood cells have microbial inclusions. The sample belonged to a person who had recently returned from a mosquito-infested region of Africa.

c. A large organism (over 3 m), found on the beach, brown in color, which stores food as mannitol.

d. A water sample from a birdbath. The numerous small organisms seen are covered with fine hairs.

e. A marine sample containing a number of small organisms with nearly symmetrical shells. Further analysis indicates that the shells are made of silicon dioxide and that the organisms are rich in oil.

39

40

Case Study Exercise 3  A Survey of Protists

f. An organism possessing lobe-shaped pseudopods, isolated from the feces of a person recently deceased after a gastrointestinal illness.

g. A seawater sample sent to your lab from the Gulf Coast. The water has an orange tint to it, and organisms in the jar have a pair of flagella and seem to rotate as they move through the water.

h. A pond water sample in which the flagellated organisms are marked by a single red spot at one end.

CASE STUDY The following incident illustrates the potential danger posed by some protista. Study the details of the case and use your knowledge of protist biology to answer the case study analysis questions.. Primary Amebic Encephalitis—Louisiana, 2011 Primary amebic encephalitis (PAM) is caused by infection with Naegleria fowleri, a microbe sometimes known as the “brain-eating amoeba” because it causes widespread destruction of brain tissue. Generally found in warm bodies of freshwater such as lakes, rivers, or hot springs, the amoeba enters the body through the nose when people are swimming, diving, or ducking under water. After entering the nose, it migrates to the brain along the olfactory nerve, initially producing symptoms similar to those of meningitis— headache, fever, nausea, vomiting—but soon worsening to include confusion, loss of balance, seizures, and hallucinations. Death is a virtual certainty (only one person has ever been thought to survive) and occurs within a week. Most infections in the United States occur in southern states during summer months where prolonged hot weather warms recreational lakes and rivers. Although N. fowleri is a fairly common inhabitant of such warm freshwater bodies, PAM is an exceedingly rare disease, with only 143 cases seen from 1962 to 2016. In June 2011, the Louisiana Department of Health and Hospitals received reports of two people, a 20-year-old man and a 51-year-old woman, dying of the

disease just a few months apart. Causing further concern was the fact that neither victim could be linked to the warm water springs, lakes, or rivers that serve as a breeding ground for the amoeba. As an investigation of both cases began, health officials worried that they could be dealing with a strain of N. fowleri that had become more virulent or that had found a new way to enter the body. After a fivemonth investigation, the Louisiana Department of Health and Hospitals issued a warning concerning the use of neti pots and infection with Naegleria fowleri. A neti pot resembles a small teapot with a long spout and is used by some people to rinse the nasal passages with a saline solution as a treatment for colds and allergies, congestion, or moistening dry nasal passages. Both Louisiana residents used neti pots filled with warm tap water to rinse their sinuses. Investigators collected water samples from both homes, and testing revealed the presence of Naegleria fowleri in each case. These two incidents represented the first time that PAM was associated with the presence of N. fowleri in household plumbing served by treated municipal water supplies, as well as the first time that infection with the amoeba was linked to use of a neti pot or similar nasal irrigation device. The Centers for Disease Control and the U.S. Food and Drug Administration now recommend that water used for the rinsing of sinuses be distilled, boiled, or filtered, that similarly treated water be used to rinse the neti pot, and that the pot be allowed to air-dry completely between uses.



Case Study Exercise 3  A Survey of Protists

41

CASE STUDY ANALYSIS 1. Naegleria fowleri is a thermophilic organism. Using the glossary of your book, explain what a thermophile is and how this characteristic relates to the epidemiology of N. fowleri infection.

2. The Centers for Disease Control have made several recommendations to reduce the likelihood of infection with N. fowleri while participating in water activities. Aside from prohibiting all swimming, what do you think they recommended?

3. Entamoeba histolytica is an amoeba responsible for the gastrointestinal disease amebic dysentery. While E. histolytica is found in the environment as both an active trophozoite and as an inactive cyst, virtually all cases of amebic dysentery can be traced to infection with the cyst form of the organism. Speculate on why the active form of the pathogen causes far fewer cases of disease.

THERE’S MORE TO THE STORY . . .

SHIPPING INFORMATION

to weekly or biweekly sampling of the same site for So, think you’re an expert on algal identification? Feel at least one year, although if your instructor has an like you could do this for a living? How about as a established sampling site, you may be able to work as volunteer? Here’s your chance. The National Oceanic part of a team for a shorter period of time. Volunteerand Atmospheric Administration (NOAA) is a federal ing begins with a pair agency that focuses on of training courses (inthe oceans and atmoperson or online) that sphere, making weather HAB SCREENING DATA SHEET together last about five predictions, tracking ATLANTIC REGION 3 hours and will ensure hurricanes, and monitorFL East Coast, GA, SC, NC South of Cape Lookout that you can identify ing fisheries. It oversees TARGET SPECIES SCREENING LIST FIELD DATA the important algal spethe National Centers for REQUIRED No Yes Elevated cies in your area. Once Coastal Ocean Science Name: Akashiwo sanguinea Alexandrium monilatum trained, you spend a (NCCOS), which in turn Ceratium furca few minutes every other run the Phytoplankton Sampling Site: Chaetoceros spp. week collecting water Monitoring Network Sample Date: Cochlodinium spp. Dinophysis spp. samples and 20 minutes (PMN), a large group Karenia spp. Sample Time: to two hours identifying of volunteer scientists Karlodinium spp. Lingulodinium polyedrum the algae present (the who collect samples of Water Temp (ºC): Prorocentrum spp. time required for idensea water, identify the Salinity (ppt): Pseudo-nitzaschia spp. Pyrodinium bahamense tification typically gets algal species present, Other Elevoted/Bloom Species shorter as you become and forward the results more experienced). to the PMN, promoting OPTIONAL Your instructor a better understanding None Present Abundant Bloom Weather: Sunny | Partly Cloudy | Centric Diatoms may have you—in addiof harmful algal blooms. Mostly Cloudy | Cloudy | Rain Pennate Diatoms tion to reporting your This is your chance to be Wind direction: N | NE | E | SE | Dinoflagellates S | SW | W | NW Cyanobacteria findings to the PMN— a primary investigator in Ciliates Wind speed (mph): 0-5 | 5-10 | record your data in a lab a real-life research proj- 10-15 | 15-20 | 20-25 | 25+ Other Zooplankton notebook and/or save ect, a rare opportunity Tides: High | Low | Incoming | Outgoing - SHIP 3 SAMPLES VIA UPS NEXT DAY AIR: 1L live whole water, 30mL preserved your sampling records, not usually available to Air Temp (ºC) whole water & 125mL preserved net tow - SHIP 2 SAMPLES VIA UPS GROUND: like the one shown here, undergraduate students. pH: Follow Shipping Calendar 30mL preserved whole water over the course of the As a PMN volunteer, Dissolved Oxygen (ppm): & 125 mL preserved net tow - SHIP 1 SAMPLE VIA UPS GROUND: semester. You can then you may also have the Barometric pressure (mmHg): Follow Shipping Calendar 30mL preserved whole water graph changes in algal opportunity to collabo- Secchi Disk (cm): - No samples needed populations as a funcrate with NOAA laboJeff Paternoster NOAA PMN, 219 Fort Johnson Rd, Charleston, SC 29412 tion of time of year, ratories and tour their 843-762-8657 | [email protected] water temperature, pholand- and sea-based toperiod (amount of daylight per 24 hours) or any facilities. other factor. The PMN will supply you with everything Ready to get started? Check out the Phytoplankyou need except for a light microscope and comton Monitoring Network at https://products.coast puter (needed for training and to enter data into the -alscience.noaa.gov/pmn/ search PMN. PMN online database). As a volunteer, you commit

REFERENCE State of Louisiana Department of Health and Hospitals. 2011. North Louisiana Woman Dies from Rare Ameba Infection. http://new.dhh.louisiana.gov /index.cfm/newsroom/detail/2332.

C A S E S T U DY E X E R C I S E

4

A Survey of Fungi STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Describe the major groups of fungi. 2. Differentiate fungal samples based on macroscopic and microscopic characteristics.

INTRODUCTION The fungi are a large group of nonmotile, eukaryotic organisms that can be found in a variety of forms, including mushrooms, puffballs, molds, and yeasts, all of which appear very different but share many biological features. All members are ­heterotrophic, meaning they require an organic source of carbon, as opposed to autotrophic (photosynthetic) organisms that can utilize CO2 from the atmosphere. Most fungi are described as saprobes, meaning they derive their energy from the decomposition of dead organisms; in fact, along with the bacteria, fungi are the primary decomposers of organic compounds on earth. This decomposition allows for the reuse of carbon and nitrogen by other organisms and prevents the planet from being swamped with organic waste. A relatively small number of fungi grow as parasites and are responsible for diseases ranging from the mundane (athlete’s foot) to the life threatening (fungal meningitis). The Greek prefix myco means fungus and helps to explain the fact that the study of fungus is mycology and a person who studies fungi is a mycologist. The common characteristics of fungi provide ample means to differentiate them from plants and animals. Unlike plants, fungi are unable to photosynthesize and contain the polysaccharide chitin in their cell walls, rather than cellulose that is commonly seen in plants. Unlike animals, fungi must digest their food before ingesting it. By secreting digestive enzymes, they can digest organic material in the surrounding environment. The simple nutrients that result are then imported into the cell. Fungi can be found in a variety of forms, many of which should already be familiar. The larger mushrooms, puffballs, and truffles (Figure  4.1) are sometimes referred to as ­macrofungi, while microscopic fungi are referred to as yeasts or molds, depending on their structure. Yeasts are single-celled fungi, with a spherical to oval shape and a typical diameter of 3–5 µm. The most common means of reproduction in yeasts is

budding, in which a new, small cell forms from an older larger one. When a string of these new cells forms, the result is a pseudohypha (Figure 4.2a). Two common yeasts with human importance are Saccharomyces cerevisiae, which is used in the manufacturing of bread and beer, and Candida albicans, part of the normal flora of the respiratory, gastrointestinal, and female reproductive tract. Overgrowth of Candida can lead to conditions such as thrush and vulva vaginitis (inflammation of the vulva or vagina) in otherwise healthy persons while more serious systemic infections may be seen in patients who are immunocompromised, diabetic, or have indwelling medical devices such as catheters or shunts. When fungi grow as strings of long, filamentous cells, they are referred to as molds. A single filament is known as a hypha (plural, hyphae) while a large intermeshed mat of hypha is called a mycelium (Figure  4.3). Hypha can be categorized as septate if individual cells in the hypha are separated by walls or as nonseptate if the hypha is continuous (Figure 4.4). Each hypha in a mycelium is capable of digesting and absorbing nutrients, making a fungus almost perfectly adapted for eating and ensuring that fungi are found in nearly every habitat on earth. Some fungi are referred to as dimorphic, capable of growing as yeasts or molds, depending on the environmental conditions. Many fungal pathogens grow in the environment as molds and then convert to a yeast form within the body. Although spores, hyphal elements, and yeast may all initiate a fungal infection upon entering the body, reproductive

Figure 4.1  Most fungi, like these mushrooms growing on a fallen log, are saprobes, gaining nutrition through the slow decomposition of organic matter. ©BadZTuA/iStock/Getty Images

43

44

Case Study Exercise 4  A Survey of Fungi Bud

Nucleus

Bud scars

Pseudohypha (a) (a)

(b)

Figure 4.2  (a) As a yeast cell reproduces, it creates buds, eventually resulting in the formation of a pseudohypha. (b) Cells of baker’s yeast (Saccharomyces cerevisciae) display many small buds and bud scars, along with a few short pseudohyphae (1200x). (b) ©Science Photo Library/ Alamy Stock Photo

(b)

spores, by virtue of sheer number, are most commonly responsible. Most respiratory mycoses (fungal infections) can be traced to the inhalation of fungal spores, while cutaneous infections begin when the body comes in contact with surfaces contaminated with spores. As an example, tinea pedis, the opportunistic

Figure 4.3  (a) To the naked eye, the mycelium of a fungus appears as a cottony interweaving of individual hyphae. (b) Scanning electron micrograph of a mycelium reveals the structure of the hyphae (x5000). (a) ©Udomsook/iStock/Getty Images; (b) ©Dr. Judy A Murphy, Microscopy & Imaging Consultant, Stockton, CA

Septa

As in Penicillium

Septate hyphae

Nonseptate hyphae

Septum with pores Nucleus

Nuclei As in Rhizopus

Figure 4.4  Fungal hyphae may be septate or nonseptate, and this characteristic can be used in the laboratory to help identify an unknown fungal species.



Case Study Exercise 4  A Survey of Fungi

45

Sporangia

Columella

(a) Mucor

Phialide

Sporangiospores (Within sporangia)

Syncephalastrum

(b) Penicillium

Phialospores (Conidia on phialides)

Gliocadium

Conidiophore

Cladosporium (c)

Blastoconidia (Formed by budding)

Oospora (d) Arthrospores (By separation)

Candida albicans (e) Chlamydospores (Large, round)

Fusarium

Alternaria Macroconidia (Multicelled conidia)

(f)

Microsporum canis

Figure 4.5  (a) Sporangiospores are asexual fungal spores borne within a dedicated structure known as a sporangium, while spores that form on specialized hyphae are called conidia. (b) Phialospores sit atop a pear-shaped cell called a phialide. (c) Blastoconidia are formed by budding from preexisting conidia. (d) Arthrospores form from preexisting hyphal cells. (e) Chlamydospores are large, thick-walled structures commonly found on older cultures of many fungi. (f) Any multicellular conidia are known as macroconidia.

mycosis more commonly known as athlete’s foot, has little to do with being an athlete but rather is connected to walking barefoot in damp locker rooms where fungi thrive. M ­ ycoses due to true fungal pathogens are thankfully rare among people with a competent immune system; however, mold spores are a common allergen, with symptoms often becoming especially pronounced when the weather is damp. Fungal reproduction is complex, with both sexual and asexual methods of reproduction being used by most species. Asexual reproduction occurs when haploid spores are released from a single parent. If these spores are contained within a saclike container called a sporangium, the spores are referred to as sporangiospores. Conidia are spores that are not enclosed within a sporebearing sac but rather form from a special fertile hyphae called a conidiophore. Both types of spores are seen in Figure 4.5. Asexual spores give rise to new fungi that are exactly like the parent that produced them. Sexual spore formation is more complex but offers the promise of more variability in the offspring. By combining genes from two parents, the offspring produced will vary in form and function both from each other and from both parents. Fungi are more likely to undergo sexual reproduction when environmental conditions are poor, for instance too warm or cold, or when there is a lack of nutrients this variability provides at least some of the offspring with advantageous traits and increases the overall survival of the species. Three types of fungal spores are seen in fungi. These are illustrated in Figure 4.6. Morphological characteristics such as the formation of spores is commonly used in the laboratory to identify and

classify fungi, even as molecular techniques play a greater role in the process. Like the protists, the fungi have never found a permanent taxonomic home of their own, bouncing from one Kingdom to another. The most current molecular evidence places them in the Supergroup Opisthokonts, which also contains the Animal Kingdom. The fungi are further divided into five major groups (Figure 4.7). Chytrids  These represent the simplest fungi. Unique to this group is the formation of a motile zoospore with a single whiplash flagellum. Most members of this group are saprobes, helping to decay dead plants and animals. Zygomycetes  This group of roughly 1000 species produces sexual zygospores when hyphae of two opposite mating strains fuse to create a diploid zygote (Figure 4.8). This zygote will eventually germinate, giving rise to a sporangium filled with sexual spores. The common bread mold Rhizopus belongs to the Zygomycetes. A cursory look

Asci Zygospore

Zygospore

Basidium

Ascospores

Figure 4.6  Types of sexual spores seen in fungi.

Basidiospores

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Case Study Exercise 4  A Survey of Fungi

Supergroup Opisthokonts

Chytrids

Animals

Fungi

Zygomycetes

Glomeromycetes

Ascomycetes

Basidiomcetes

Figure 4.7  Taxonomic organization of fungi.

at a Rhizopus colony can give a clue to its age; the whitish mycelium will grow darker as dark sporangia develop (Figure 4.9). Glomeromycetes  The 200 or so species in this group almost all exist in a mutualistic relationship with plant roots, forming a structure called an arbuscular mycorrhizae. Ascomycetes  Spores formed within a fungal sac, or ascus, are the hallmark of this large group (65,000 species). The ascus develops from the union of two hyphae of opposite mating type. Meiosis, sometimes followed by mitosis, results in the formation of four to eight haploid ascospores (Figure 4.10). Many species of great importance to humans are members of the Ascomycetes. Species of Aspergillus can be found growing on fruit and bread, and industrial uses of the fungus include the production of citric acid and soy sauce. Some species of Aspergillus are opportunistic pathogens and are responsible for aspergillosis, a collection of mostly respiratory diseases of varying severity. Histoplasma capsulatum is associated with bird and bat droppings and causes a respiratory infection known as Ohio Valley fever, while members of the genus Trichophyton are responsible

Figure 4.8  The sexual zygospore of Rhizopus forms between two hyphae of opposite mating strains (300x). ©McGraw-Hill Education/Richard Gross, photographer

Figure 4.9  Several species of fungi growing on bread. The darker areas of each colony represent areas of sporangia formation. ©imageBROKER/Alamy Stock Photo

Figure 4.10  The cup fungus Peziza produces eight ascospores in each saclike sporangium. ©McGraw-Hill Education/Richard Gross, photographer



Case Study Exercise 4  A Survey of Fungi

Figure 4.11  Onychomycosis (nail infection) caused by Trichophyton

Basidiomycetes  Basidium refers to the small clublike structure that supports the spores of fungi in this group (Figure 4.13). Most common mushrooms, puffballs, and bracket fungi are classified here. The basidiospores are found attached to basidia, which in turn are located within the gills of the mushroom. Many members of the ascomycetes and basidiomycetes have no known sexual stage in their life cycle. Because this posed problems for a taxonomic system that classified fungi by the type of sexual spores they produced, mycologists created an artificial group, the Deuteromycota, or fungi imperfecta. This group consisted of fungi with no known sexual stage. If a sexual stage was discovered for a particular fungus, the fungus was reclassified into its proper group (ascomycetes, basidiomycetes, etc.). The problem with this process, however, is that a fungus can be known by two completely different names, both of which are correct. One way of

rubrum, which displays an affinity for skin, hair, and nails. Trichophyton infections of the skin are referred to variously as ringworm, athlete’s foot, jock itch, or tinea. Source: CDC/Dr Edwin P Ewing, Jr.

for superficial fungal infections of the skin, nails, and hair (Figure 4.11). Other notable members of the phylum include the baker’s yeast Saccharomyces cerevisiae and Penicillium (Figure 4.12), species of which are used to produce the antibiotic penicillin as well as Roquefort and Camembert cheeses.

(a)

(b)

Figure 4.12  Colonies of Penicillium (1x) growing on an agar plate. Inset displays the conidiophore, with two brush-shaped clusters of spores (970x). (a) ©grebcha/iStock/Getty Images; (b) Source: CDC/Dr. Lucille K. Georg

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Figure 4.13  (a) The gills of a wild mushroom. Mushrooms are the fruiting bodies of a fungus, and the basidia (bearing basidiospores) are attached to the gills on the underside of the mushroom cap. (b) Basidiospores on the gill of a Coprinus mushroom (430x). (a) ©IT Stock/age fotostock; (b) ©McGraw-Hill Education/Richard Gross, photographer

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Case Study Exercise 4  A Survey of Fungi

thinking of this is to consider a person who takes the name of his or her spouse after marriage; both names refer to the same person, but until this fact is known, some confusion will no doubt ensue. Genetic analysis is slowly alleviating this problem, as fungi are more often classified by their DNA rather than by their physical appearance. Until genetic analysis becomes routine, initial classification of a fungus is typically accomplished through visual examination of a sample. First, the color, texture, and morphology of a mold colony, both top and bottom, is examined. The texture of a fungal colony

can be described as leathery, velvety, cottony, or powdery, while common morphologies include flat, verrucose (rough, hilly), and cerebriform (brainlike). Color is self-explanatory with the caveat that the color of a colony generally will change as it ages and forms sporangia. After the initial macroscopic assessment, microscopic examination of hyphae (septate or not), sporangia, and spores can be used to complete the initial identification (Figure 4.14). The microscopic appearance of a number of common fungi is seen in Figure 4.15, while the fungus gallery (Figure 4.16) contains photographs of a variety of fungi.

(a)

(b)

(c)

Figure 4.14  (a) A colony of Trichophyton rubrum is velvety, cerebriform, and a yellow-beige on the upper surface. (b) The same colony is a yellow to brown color on the reverse. (c) A microscopic view of Trichophyton (1000x) reveals septate hyphae and numerous microconidia attached directly to the hyphae. (a-b) Source: CDC/Dr. Libero Ajello; (c) Source: CDC



Case Study Exercise 4  A Survey of Fungi

(1)

(2)

(6)

(11)

(7)

(12)

(16)

(3)

(17)

(4)

(8)

(13)

(18)

(1) Penicillium– bluish-green; brush arrangement of phialospores. (2) Aspergillus– bluish-green with sulfur-yellow areas on the surface. Aspergillus niger is black. (3) Verticillium– pinkish-brown, elliptical microconidia. (4) Trichoderma– green, resemble Penicillium macroscopically. (5) Gliocadium– dark-green; conidia (phialospores) borne on phialides, similar to Penicillium; grows faster than Penicillium. (6) Cladosporium (Hormodendrum)– light-green to grayish surface; gray to black back surface; blastoconidia. (7) Pleospora– tan to green surface with brown to black back; ascospores shown are produced in sacs borne within brown, flask-shaped fruiting bodies called pseudothecia. (8) Scopulariopsis– light-brown; rough-walled microconidia. (9) Paecilomyces– yellowish-brown; elliptical microconidia. (10) Alternaria– dark greenish-black surface with gray periphery; black on reverse side; chains of macroconidia. (11) Bipolaris– black surface with grayish periphery; macroconidia shown.

Figure 4.15  Microscopic appearance of some common molds.

(9)

(14)

(19)

(5)

(10)

(15)

(20)

(12) Pullularia– black, shiny, leathery surface; thick-walled; budding spores. (13) Diplosporium– buff-colored wooly surface; reverse side has red center surrounded by brown. (14) Oospora (Geotrichum)– buff-colored surface; hyphae break up into thin-walled rectangular arthrospores. (15) Fusarium– variants, of yellow, orange, red, and purple colonies; sickle-shaped macroconidia. (16) Trichothecium– white to pink surface; two-celled conidia. (17) Mucor – a zygomycete; sporangia with a slimy texture; spores with dark pigment. (18) Rhizopus– a zygomycete; spores with dark pigment. (19) Syncephalastrum– a zygomycete; sporangiophores bear rod-shaped sporangioles, each containing a row of spherical spores. (20) Nigrospora– conidia black, globose, one-celled, borne on a flattened, colorless vesicle at the end of a conidiophore.

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(a) Rhizopus sporangia (430x). A zygomycetes, Rhizopus commonly colonizes bread, where it produces light green to black colonies.

(b) Pilobolus sporangia (25x). Pilobolus spores are eaten by herbivores as they graze. The spores survive the gastrointestinal tract and are ultimately deposited in manure. As the sporangia grow, they orient themselves toward light and eventually burst, casting spores as far as 2 meters away.

(c) A fairy ring. The mushrooms erupting from the grass indicate the spread of the fungal mycelium below the surface of the soil. Folklore has traditionally associated fairy rings with bad luck or danger. (d) Black truffles (Tuber melanosporum) grow a few cm below ground in a symbiotic relationship with the roots of certain plants. Some breeds of dogs and pigs have a natural ability to smell the chemicals exuded by truffles growing as much as a meter below the surface. Truffles like these sell for hundreds of dollars per pound.

(e) Botrytis is a plant pathogen. One species grows on wine grapes, giving the wine a distinct, sweet taste. Grapes covered in Botrytis are said to possess the “Noble Rot”.

Figure 4.16  Fungus gallery. (a,b) ©McGraw-Hill Education/Richard Gross, photographer; (c) ©NajaShots/iStock/Getty Images; (d) ©bonzami emmanuelle/Alamy Stock Photo; (e) Source: CDC/Dr. Lucille K. Georg; (f) Source: CDC; (g) Source: CDC/ Dr. Libero Ajello; (h) Source: CDC; (i) Source: CDC; ( j) Source: CDC/Dr. Lucille K. Georg; (k) Source: CDC/Dr. Libero Ajello; (l-1) ©Bear Dancer Studios/Mark Dierker; (l-2) ©Worraket/Shutterstock; (l-3) ©Jorgen Bausager/Getty Images

(f) Paracoccidioides brasiliensis during its yeast phase. Found in the rich soils of Central and South America, Paracoccidioides is one of the four genera of true fungal pathogens. Inhalation of conidia can lead to paracoccidioidomycosis. Yeast colonies are described as white, heaped, wrinkled or folded and appear only after incubation at 37°C for 10–20 days.

(g) Microscopic morphology of Histoplasma capsulatum (400x). Found growing in soils containing high concentrations of bird and bat guano in the southern and eastern United States, as well as portions of South America and Africa. Inhalation of conidia can lead to histoplasmosis—also called Ohio Valley fever—a disease typically confined to the lungs but which can be lethal if other organs are affected.

(i) Sporothrix schenckii (500x). A member of the ascomycetes, S. schenkii can cause sporotrichosis, or rose handler’s disease.

(k) Frontal view of a Madurella grisea colony. A member of the ascomycetes, M. grisea is found in soil in tropical and subtropical areas of Africa, India, and South America. The fungus enters the body via trauma, often to the foot, and causes slow progressing infections known as mycetomas that are characterized by large black masses of hyphae.

(h) Oral thrush caused by Candida albicans. Found primarily in babies and older adults, whose weaker immune systems are not strong enough to limit growth of the yeast.

( j) Aspergillus niger (500x). Aspergillus is a common opportunistic fungus that may infect individuals whose immune systems have been compromised. A member of the ascomycetes, it displays septate hyphae and long conidiophores that terminate in rough, darkly colored conidia.

(l) Mushrooms. The fruiting bodies of basidomycetes, mushrooms display a variety of morphological types.

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Case Study Exercise 4  A Survey of Fungi

PRE-LAB QUESTIONS Remember, Understand, Apply 1. All fungi are a. photosynthetic. b. heterotrophic. c. mixotrophic. d. autotrophic. 2. Fungi that decompose dead organic material are said to be a. parasites. b. anaerobes. c. colonial. d. saprobes. 3. Which Greek prefix indicates fungi? a. myco b. auto c. sapro d. pseudo 4. Fungi a. typically reproduce sexually when environmental conditions are poor. b. typically reproduce sexually when environmental conditions are good. c. always reproduce sexually. d. always reproduce asexually. 5. Yeasts form hyphae while molds form pseudohyphae. a. true b. false 6. Molds are made up of _____ which, when grouped together are referred to as _____. a. buds, spores b. hyphae, mycelium c. hyphae, sporangia d. pseudohyphae, conidia 7. What word describes a hypha that is divided into small sections by crosswalls? a. sexual b. biphasic c. conidial d. septate 8. Which fungal group possesses motile spores? a. chytrids b. zygomycetes c. glomerulomycetes d. basidiomycetes 9. Which characteristic is generally not used in the initial identification of a fungus? a. appearance of the mycelium b. type of sporangia produced

c. food source d. structure of the hyphae 10. Most mushrooms are categorized within the group a. chytrids. b. zygomycetes. c. glomerulomycetes. d. basidiomycetes.

MATERIALS Each group should obtain: Sealed Sabouraud dextrose agar plate cultures of: Rhizopus sp. Penicillium sp. Aspergillus sp. These plates should be sealed with parafilm or tape. Do not remove the lids from the plates because the number of spores released could constitute a hazard. Sabouraud dextrose agar plate culture of: Saccharomyces cerevisiae Prepared slides of: Aspergillus sp. Rhizopus sp. Penicillium sp. Methylene blue, iodine, or lactophenol cotton blue

PROCEDURE Saccharomyces cerevisiae 1. Prepare a wet mount of Saccharomyces cerevisiae. The procedure for preparing a wet mount may be reviewed in Exercise 3. ∙∙ Begin by adding a single drop of dye to the center of the slide. ∙∙ Use an inoculating needle to aseptically add a small amount of yeast to the dye. ∙∙ Gently place a cover slip atop the dye. 2. Examine the slide using low power. When you find an area of great interest, swing your high-power objective into position. Sketch a portion of the microscopic field, looking especially for budding cells and pseudohyphae. Rhizopus sp. 1. Examine a colony of Rhizopus macroscopically. Do not remove the cover from the plate. Sketch the colony, and provide a description of the color (both top and bottom of the colony), texture (leathery, velvety, cottony, or powdery), and surface morphology (flat, verrucose, or cerebriform) as described in the Introduction.



Case Study Exercise 4  A Survey of Fungi

2. Using a dissecting microscope, examine the colony and sketch the details of the mycelium, indicating hyphae and sporangia. 3. Using a compound microscope, examine prepared slides of Rhizopus. Sketch a portion of the microscopic field, being certain to identify hyphae, sporangia, and spores. Label each of these structures. Penicillium sp. 1. Examine a colony of Penicillium macroscopically. Do not remove the cover from the plate. Sketch the colony, and provide a description of the color (both top and bottom of the colony), texture (leathery, velvety, cottony, or powdery) and surface morphology (flat, verrucose, or cerebriform) as described in the Introduction. 2. Using a dissecting microscope, examine the colony and sketch the details of the mycelium, indicating hyphae and conidia. 3. Using a compound microscope, examine prepared slides of Penicillium. Sketch a portion of the microscopic field, being certain to identify hyphae, conidiophores, and conidia. Label each of these structures.

53

Penicillium is often grown on oranges, which are then used to prepare slides. Find the edge of the orange peel under low power; the mold will be visible on the surface, looking almost like small plants erupting from the Earth.

Aspergillis 1. Examine a colony of Aspergillis macroscopically. Do not remove the cover from the plate. Sketch the colony, and provide a description of the color (both top and bottom of the colony), texture (leathery, velvety, cottony, or powdery) and surface morphology (flat, verrucose, or cerebriform) as described in the Introduction. 2. Using a dissecting microscope, examine the colony and sketch the details of the mycelium, indicating hyphae and conidia. 3. Using a compound microscope, examine prepared slides of Aspergillis. Sketch a portion of the microscopic field, being certain to identify hyphae, conidiophores, and conidia. Label each of these structures.

RESULTS Record your results. For each sample, provide a drawing that indicates the major features seen in both macroscopic and microscopic views of the organism. Also indicate the total magnification used to observe the specimen, whether it is 1x (naked eye), 20x or 40x (dissecting microscope), or 100x or 450x (compound microscope), as well as any colony morphology. S. cerevisiae (______x)

Rhizopus sp. (______x)

Rhizopus sp. (______x)

Rhizopus sp. (______x)

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Case Study Exercise 4  A Survey of Fungi

Penicillium sp. ( x)

Penicillium sp. ( x)

Penicillium sp. ( x)

Aspergillus sp. ( x)

Aspergillus sp. ( x)

Aspergillus sp. ( x)



Case Study Exercise 4  A Survey of Fungi

REVIEW QUESTIONS Analyze, Evaluate, Create 1. Differentiate between a hypha, a pseudohypha, and a mycelium.

2. Sketch an example of septate hypha and nonseptate hypha.

3. What is a mycosis?

4. Describe and draw the principal differences between a yeast and a mold.

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Case Study Exercise 4  A Survey of Fungi

5. Briefly, what are the advantages and disadvantages of sexual reproduction in fungi?

6. Briefly, what are the major characteristics used to differentiate a. Fungi from plants?

b. Fungi from animals?

c. Deuteromycota from each of the other fungal groups?



Case Study Exercise 4  A Survey of Fungi

57

CASE STUDY While not as common as those due to bacteria or viruses, serious fungal infections certainly occur, either alone or as part of a larger outbreak. Study the following case and use your knowledge of fungi to answer the case study analysis questions.

source of the outbreak in order to prevent further infections. Initial testing revealed 17 counselors (53%) whose blood serum or urine contained antigens from Histoplasma capsulatum; these were deemed confirmed cases of the disease. Of 153 responses to a questionnaire sent to camp attendees, Histoplasmosis Outbreak among Day Camp Attendees— 17 campers (11%) had confirmed or suspected histoplasmosis Nebraska, June 2012 (suspected cases were those in which illness was present but Mention microbial infections and most people think of hospilaboratory results were inconclusive or unavailable). tals, not summer camp. But being outdoors in the sunshine and Prior to the beginning of camp, the counselors participated fresh air is not without risk. In June 2012, a camp counselor in in a camp cleanup, where the 12 campsites were prepared for Omaha, Nebraska, was diagthe summer. Preparations nosed with histoplasmosis, a included raking leaves, cleanfungal infection with symping picnic tables, digging fire pits, and moving firewood. toms similar to p ­neumonia; Counselors reported that they fever, headache, and respifound bird and bat guano on ratory distress. Although picnic tables and the dirt floor most people suffering from of camp shelters at two of the histoplasmosis will recover campsites. Campers arrived without medical treatment, a week later and took part in the very young, very old, and nature walks, outdoor games, people with chronic lung conwilderness training, and arts ditions are at greater risk for and crafts, but did not engage developing severe disease, in high-risk activities like which may on rare occasions those performed by the counprove fatal. Histoplasmosis is selors earlier in the summer. caused by inhaling spores of Macroconidia of Histoplasma capsulatum. Macroconidia of Histoplasma capsulatum. Source: CDC/Dr. Libero Ajello Analysis of data collected the fungus Histoplasma capby the DCDH and NDHHS sulatum, which is common in determined that campers assigned to campsites where guano the United States, especially in the Mississippi and Ohio River was seen were 2.4 times more likely to become ill than those valleys. The spores are typically present in dirt contaminated campers assigned to campsites greater than 20 yards away. with bird or bat guano and activities like renovation of old In fact, the data showed that the further away campers were buildings, cave exploration, and even gardening, are known to from a site where guano was found, the less likely they were increase the risk of infection. to become ill. Based on this report, the camp was relocated Because the sick camp counselor was one of 32 who to a different park, and the city’s parks and recreation division worked at the camp, and the camp served 797 children, was given recommendations to prevent bat roosting, recogthe Douglas County Department of Health (DCDH) and nize potentially contaminated areas, and properly deal with the Nebraska Department of Health and Human Services such sites. (NDHHS) began an investigation to determine the extent and

CASE STUDY ANALYSIS 1. Exploring caves is a known risk factor for contracting histoplasmosis while wrestling is a known risk factor for contracting ringworm (a fungal infection of the skin). Explain why each activity increases the risk of contracting a specific fungal infection.

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2. Treatment of fungal infections generally takes much longer than treatment of bacterial infections, and the drugs used to treat these infections are generally more toxic to humans than are antibacterial drugs. Speculate on why this may be (comparing fungal, bacterial, and human cells may help).

3. Use the Internet to find the target (cellular structure or function) of each of the antifungal drugs in the left column, and match each drug to its target in the right column. Flucytosine Amphoteracin B Imidazole

a.  Cell membrane b.  Cell wall c.  Cell division d.  Nucleic acid synthesis

Nystatin Griseofulvin Echinocandins Terbinafine

REFERENCES CDC. Histoplasmosis. http://www.cdc.gov/nczved/divisions/dfbmd/diseases/histoplasmosis. CDC 2012. Histoplasmosis outbreak among day camp attendees—Nebraska, June, 2012. Morbidity and Mortality Weekly Report, 61 (37): 747–748.

C A S E S T U DY E X E R C I S E

5

A Survey of Parasitic Worms STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Understand how helminthic parasites are identified. 2. Use visual means to differentiate a number of parasitic worms.

INTRODUCTION The study of worms that parasitize humans offers an entirely different experience from that of studying prokaryotic or even small eukaryotic parasites. Helminths (parasitic worms) are comparatively large, multicellular animals with tissues and organ systems. Their biology is in most cases far closer to the humans they parasitize than to other microbes, a fact that makes effective treatment of helminthic infections difficult. Epidemiologists estimate that about 1.5 billion people are infected with helminths worldwide, mostly confined to

Earth’s tropical regions. In the laboratory, helminths are easily distributed among three categories through their physical appearance: nematodes (roundworms), cestodes (flatworms, or tapeworms), and trematodes (flukes). These groups can be seen in Figure 5.1. Although the biology of the organisms in each category is complex, with complicated life cycles, multiple hosts, and various levels of pathogenicity, diagnosis of infection generally relies on the visualization of the parasites, as part of an O and P—Ova and Parasite— test. In such a test, a stool sample is examined under a microscope for the presence of eggs (ova) or worms (parasites). A skilled parasitologist is often able to identify the organism responsible for an infection simply by examining the eggs in the feces, without ever having to visualize the worm itself. This fact helps to explain the importance of helminth egg identification as a diagnostic tool.

(b)

(a)

Figure 5.1  Parasitic worms are divided into three broad categories by shape. (a) Nematodes, or roundworms (Ascaris lumbricoides). (b) Cestodes, or tapeworms (Taenia saginata). (c) Trematodes, or flukes (Fasciola hepatica). (a) ©McGraw-Hill Education/Lisa Burgess, photographer; (b) Source: CDC; (c) ©D. Kucharski K. Kucharska/Shutterstock

(c)

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Case Study Exercise 5  A Survey of Parasitic Worms

(a) (a)

(b)

Figure 5.3  Strongyloides. (a) Strongyloides (100x) filariform larvae. The Strongyloides nematode has a complex life cycle that alternates between free-living (rhabditiform) and parasitic (filariform) cycles. (b) Mouthparts of Strongyloides rhabditiform larva (1000x). The presence of rhabditiform larvae in the stool is diagnostic of an infection. (a) Source: CDC - PHIL; (b) Source: CDC (b)

Figure 5.2  Hookworm. (a) Hookworm filariform larvae (100x). Two species of hookworm, Ancylostoma duodenale and Necator americanus, infect humans. As part of their life cycle, both species pass through an infectious filariform stage and a noninfectious rhabditiform stage. (b) Mouthparts of hookworm rhabditiform stage (1000x). Microscopic identification of eggs in the stool is the most common method for diagnosing hookworm infection. (a) Source: CDC - PHIL; (b) Source: CDC

Nematodes The nematodes (roundworms) are a large group thought to number as many as 1,000,000 species. The smallest nematodes are truly microscopic, while others range from just barely visible (most) to over a meter in length (very few). Hookworms (Figure 5.2) and

Strongyloides (Figure 5.3), are known as soil-transmitted helminths because their eggs hatch in soil, releasing immature worms (rhabditiform larvae). Filariform larvae can pass directly through human skin, and walking barefoot on contaminated soil is the primary means of infection. Once in the body, larvae mature into adults within the small intestine, and eggs are passed in the stool, completing the life cycle. Over 1 billion persons are infected with the soil-transmitted helminth Ascaris lumbricoides (Figure 5.4), and an additional 800 million with whipworm (Trichuris trichuria, Figure 5.5). Infection with either worm occurs when eggs are ingested, either as a result of poor handwashing or by consuming vegetables or fruits that are contaminated with eggs. Filariasis is caused by microscopic roundworms that live within the lymphatic system (Figure 5.6). The worms block the



Case Study Exercise 5  A Survey of Parasitic Worms

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(a)

(b)

(c)

Figure 5.4  Ascaris lumbricoides. (a) These nematodes (roundworms) live in the small intestine where females—the larger of the two sexes—can grow to 30 cm in length. A female may produce 200,000 eggs per day, which pass from the body along with feces, but only fertilized eggs are infectious. (b) An unfertilized egg (400x) is elongated, covered with a thick, often rough layer, and is larger than a fertilized egg. (c) A fertilized egg (400x) is rounded, thick shelled, and smaller than its unfertilized counterpart. Source: CDC

Figure 5.5  Egg of Trichuris trichiura, the causative agent of trichuriasis, or human whipworm infection. The eggs are recognized by their elongated shape, thick shell, and polar plugs. After ingestion, eggs hatch in the small intestine and release larvae, which mature into adults within the ascending colon, where they live for about a year. Sixty to 70 days after infection, the female will begin to oviposit, shedding between 3000 and 20,000 eggs per day into the feces. The presence of eggs within the feces is diagnostic of infection. Whipworm is the third most common roundworm infection of humans, with an estimated 800,000 persons infected worldwide, including in the United States. Source: CDC/Dr. Mae Melvin

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Case Study Exercise 5  A Survey of Parasitic Worms

(b)

(a)

(d)

Figure 5.6  Filarial worms. Filariasis is caused by the infiltration of small roundworms (nematodes) into the lymphatic and subcutaneous tissue. Microscopic identification of microfilariae is the most common diagnostic procedure. While eight species infect humans, most morbidity due to filariasis can be traced to just three species. (a) Wuchereria bancrofti (460x) is the most common filarial parasite in humans and is found in tropical areas throughout the world. (b) Brugia malayi (630x) causes lymphatic filariasis and is limited to Asia. (c) Infection with filarial worms can cause lymphadema, a swelling of the lower extremities. (d) Onchocerca volvulus (460x), seen here developing in its host, the black fly (Simulium ochraceum), causes onchocerciasis, or river blindness. It is found primarily in Africa, with a small area of endemicity in Latin America and the Middle East. (a, b) (c)

Source: CDC/Dr. Mae Melvin; (c) Source: Caitlin Worrell, Epidemiologist, Center for Global Health; (d) Source: CDC/Dr. R.C. Collins



Case Study Exercise 5  A Survey of Parasitic Worms

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flow of lymph, causing the testicles or lower extremities to swell, a condition known as lymphedema (or occasionally, elephantiasis). Other roundworms that infect humans include Dranunculus medinensis (Figure 5.7), Trichinella spiralis (Figure 5.8), and Enterobius vermicularis (Figure 5.9).

(a)

Figure 5.7  Dranunculus medinensis (125x), also known as Guinea worm. Humans become infected by drinking unfiltered water containing small crustaceans infected with D. medinensis. Male and female worms emerge from the crustacean and mate within the human host. After the death of the male worm, the female migrates toward the skin. About one year after infection, the female induces the formation of a painful blister on the lower extremity. When the lesion comes in contact with water (often done by the host to relieve the pain of the lesion), the female worm emerges from the body and releases larvae. Dranunculiasis has been drastically reduced as a result of intensive eradication efforts and is now restricted to rural, isolated areas in a small number of African countries. Source: CDC/Dr. Mae Melvin

(b)

Figure 5.9  Enterobius vermicularus. (a) Head region of Enterobius vermicularis (100x). Enterobiasis, or pinworm infection, is the most common helminthic infection in the United States and is especially prevalent in young children. (b) Enterobius eggs. Gravid females migrate out of the anus of an infected individual and deposit eggs on the perianal folds. Hands that have scratched the perianal area may transfer the eggs to the mouth (self-infection), while contaminated clothes, bedding, or carpeting can spread eggs to others (400x). Source: CDC

Cestodes

Figure 5.8  Trichinella spiralis cysts embedded in muscle tissue. Humans are infected by eating improperly cooked meat of animals (swine, bears, and walruses primarily) infected with the worm. Source: CDC

Tapeworms, or cestodes are named for their long ribbon-like bodies. The biology of tapeworms involves a definitive—or final— host, where adults are found, along with one or more intermediate hosts, where immature (larval) worms are found. The life cycle of cestodes typically begins when eggs are passed in the feces of a definitive host and are ingested by an intermediate host, typically another species, where the larvae develop and encyst within the tissues. When the intermediate host is eaten, larvae develop into adult worms within the small intestine, completing the life cycle. It is worth noting that tapeworms are often named for their intermediate hosts, for example, the fish tapeworm, beef tapeworm, or

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Case Study Exercise 5  A Survey of Parasitic Worms

pork tapeworm. Tapeworms possess a scolex at one end, adorned with hooks and suckers, which allows the worm to attach to the wall of the intestine. Thousands of individual segments called proglottids stretch away from the scolex, with those proglottids further from the scolex laden with eggs contained within a branching uterus. Each day, the last few proglottids break away from the worm and are passed in the feces. The presence of eggs or proglottids in the feces is diagnostic of infection. A variety of tapeworms can be seen in Figures 5.10 to 5.13.

Figure 5.12  Scolex of Taenia solium. Scolex of the pork tapeworm Taenia solium, with its characteristic two rows of hooks and four suckers, all of which allow the tapeworm to attach to the intestine and grow to a length of 7 m. The beef tapeworm Taenia saginata typically grows to 5 m or less but otherwise appears very similar to T. solium. Humans are the only definitive hosts for either species. Source: CDC

Figure 5.10  Egg of Diphyllobothrium latum (1000x). Also known as the fish tapeworm or broad tapeworm, D. latum is the largest human tapeworm, routinely growing to 10 m in length. Several intermediate hosts are involved in the life cycle, including freshwater crustaceans and minnows, but humans tend to be infected by larger freshwater fish such as trout or perch that may be eaten raw or undercooked. Mature worms reside in the small intestine where they can release more than 1,000,000 eggs per worm per day. Eggs are passed in the feces and infect the crustaceans, continuing the cycle. Source: CDC (a)

Figure 5.11  Echinococcus granulosus (45x). Echinococcus granulosus, the cestode responsible for echinococcosis—also known as hydatidosis, or hydatid disease—will grow to 3–6 mm long as an adult, with a scolex and three proglottids. Dogs and other canids are the only definitive hosts, with humans becoming infected only after ingesting food or water contaminated with dog feces. Source: CDC/Dr. Peter M. Schantz

(b)

Figure 5.13  (Continued)



Case Study Exercise 5  A Survey of Parasitic Worms

65

(c)

Figure 5.13  Gravid proglottids of Taenia saginata and Taenia solium. The presence of proglottids or eggs in the feces is diagnostic of infection, and the morphology of the proglottid is specific for each species of cestode. (a) T. saginata (8x) has 12–30 lateral uterine branches (the dark staining regions) on each side of the uterus. Adults consist of a scolex along with 1000 to 2000 proglottids, each containing about 100,000 eggs. (b) T. solium (8x) has about 1000 proglottids, each containing about 50,000 eggs. For both species, an average of six proglottids per day are passed through the stool. (c) A single Taenia sp. egg (400x). Eggs from all Taenia species are morphologically indistinguishable from each other and can survive for months in the environment. Pigs (T. solium) or cows (T. saginata) become infected by ingesting vegetation contaminated with eggs or gravid proglottids. Humans eventually become infected by eating raw or undercooked meat. Both species are found throughout the world, with T. solium being prevalent in poorer communities where humans live in close contact with pigs and are more likely to eat undercooked pork. Because the dietary restrictions of Islam and Judaism forbid the eating of pork, T. solium infection is very rare in Muslim or Jewish countries. (a-b) Source: CDC; (c) Source: CDC/Dr. Mae Melvin

Figure 5.14  Schistosoma cercaria (150x). The cercaria is the infective, free-living stage of the Schistosoma parasite. This stage is produced in the snail, which is the intermediate host of Schistosoma. After exiting the snail, the cercaria penetrates the skin of a human host, shedding its forked tail as it does so. Source: CDC/Minnesota Department of Health, R.N. Barr Library; Librarians Melissa Rethlefsen and Marie Jones, Prof. William A. Riley

Trematodes Trematodes, or flukes, are generally flattened ovoid parasites, although one group, the schistosomes, look more like cylindrical worms. Trematodes have complex life cycles, involving freshwater snails as an intermediate host. For blood flukes of the genus Schistosoma, the life cycle begins when humans urinate or defecate directly into freshwater, or when human waste is used to fertilize soil. After passing through an intermediate snail host, cercaria (Figure 5.14) are released and eventually burrow through the skin of humans, with the worm maturing in the liver and moving to the blood vessels. After mating (Figure 5.15) eggs are released into the feces, continuing the cycle. The presence of eggs in the feces is diagnostic of infection and often allows identification of the infecting species (Figure 5.16).

Figure 5.15  Schistosoma mansoni. During mating, the smaller, darker female is cradled within the gynecophoral canal of the male (100x). Source: CDC/Dr. Shirley Maddison

The life cycle of lung and liver flukes (see Figure 5.1c) begins with ingestion of the fluke, which typically occurs when eating raw watercress, undercooked fish or crustaceans, or drinking contaminated water. The presence of eggs in the feces is diagnostic of infection (Figure 5.17 and 5.18).

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Case Study Exercise 5  A Survey of Parasitic Worms

(a)

(b)

(c)

Figure 5.16  Schistosoma eggs. Eggs from trematodes responsible for schistosomiasis. Three species are responsible for the vast majority of human infection. The presence of eggs in the feces or urine, depending on the species, is indicative of an infection and aids in the identification of the species responsible. (a) Schistosoma haematobium (500x). The egg is recognized by its terminal spine. (b) Schistosoma mansomi (500x) has a lateral spine. (c) Schistosoma japonicum (500x) lacks a distinct spine. Source: CDC

Figure 5.17  Fasciola hepatica egg (125x). Humans become infected with F. hepatica by ingesting metacercariae-containing freshwater plants, especially watercress. Development of the adult fluke from the infective metacercariae takes place over 3 to 4 months in the liver, where the adult flukes reside within the biliary ducts. Human infections with F. hepatica are found in areas where sheep and cattle are raised, and where humans consume raw watercress, including Europe, the Middle East, and Asia. Microscopic identification of eggs, or rarely adult flukes, in the feces is the most common means of diagnosis. Source: CDC



Case Study Exercise 5  A Survey of Parasitic Worms

(a)

(b)

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Figure 5.18  Trematode eggs. (a) The egg of the trematode Clonorchis sinensis (400x), or Chinese liver fluke, is generally 27–35 mm in length, 11–20 mm in width, and contains an easily visible operculum at the smaller end of the egg. C. sinensis is endemic to parts of Asia, including Korea, China, Taiwan, and Vietnam. It has also been seen in nonendemic areas—including the United States—where it is invariably linked to recent immigrants or the ingestion of imported, undercooked, or pickled freshwater fish containing metacercariae of the parasite. (b) Egg of the trematode Paragonimus westermani (128x), or Oriental lung fluke, is several times larger than Clonorchis and appears asymmetrical, with one side slightly flattened. The operculum is visible on the large end while the opposite end of the egg is thickened. Human infection with P. westermani occurs by eating inadequately cooked or pickled crab or crayfish that harbor metacercariae of the parasite, which is found throughout southeast Asia and Japan. Source: CDC

PRE-LAB QUESTIONS Remember, Understand, Apply 1. Biologically, helminths are closest to a. bacteria. b. protists. c. humans. d. fungi. 2. Treatment of helminthic infections is often difficult because a. there are no drugs to treat helminth infections. b. helminths are needed by humans to digest food and extract vitamins. c. drugs which are effective at killing worms are often dangerous to their human host. d. helminths offer their hosts protection against viral infection. 3. Which of the following is not a group of parasitic worms a. roundworms. b. spirillum. c. cestodes. d. flukes.

4. Parasitic infections are often diagnosed as a result of a _____ test. a. O and P b. D and C c. Dx and Rx d. PID 5. Parasitic worm infections may be identified without ever actually seeing the worm. a. true b. false 6. Which organism is a nematode? a. Schistosoma b. Strongyloides c. Fasciola d. Taenia 7. Adult worms are found in the _____ host. a. initial b. intermediate c. secondary d. final

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8. Worms are generally named after their _____ host. a. initial b. intermediate c. definitive d. final 9. Cestodes are a. roundworms. b. tapeworms. c. flukes. d. annelids. 10. Trematodes commonly spend part of their lifecycle in a. cattle. b. pigs. c. mosquitoes. d. snails.

MATERIALS Each student should have access to the following: Prepared slides of Taenia, Ascaris, Necator, Trichinella, Facsiola, Planaria Prepared slides of Trichuris eggs Living cultures of Planaria Dissecting microscope

PROCEDURE

Planarian Wet Mount Prepare a wet mount slide of a living Planarian as follows: 1. Add a few drops of purified water (bottled drinking water is fine) to an empty Petri dish. 2. Using a Pasteur pipette, carefully draw a single planarian into the tip of the pipette. (If the planarian moves beyond the tip, it may attach to the inside of the pipette and become difficult to remove.) 3. Gently expel the planarian into the water in the Petri dish. 4. Examine the planarian, paying close attention to its movement. Record your observation in the results section. 5. Transfer the planarian to a microscope slide and gently lower a cover slip on top of it. Using the scanning and low power objectives of your microscope, study the digestive system of the planarian under low power. You should be able to see the tract split into two halves near the midpoint of the body. The light area near the center of the planarian is the pharyngeal cavity. 6. Return the planarian to the culture jar when you’ve finished your observations.

Prepared Slides Examine the prepared slides available in your lab, using the illustrations within the exercise as a guide. Record your observations in the results section.

RESULTS Record your results. For each sample, provide a drawing that indicates the major features seen in the organism. Also indicate the total magnification used to observe the specimen (i.e., 20x, 100x, 450x). ____________ (______X)

____________ (______X)

____________ (______X)

____________ (______X)



Case Study Exercise 5  A Survey of Parasitic Worms

____________ (______X)

____________ (______X)

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REVIEW QUESTIONS

Analyze, Evaluate, Create 1. Factory farming, where pigs are raised in enclosed buildings, has been disparaged as cruel, but these pigs have a lower incidence of Trichinella infection when compared to pigs that are allowed to roam free. Why, specifically, do you think this is?

2. Pinworm (Enterobius vermicularis) infection commonly spreads in day care centers. Based on the life cycle of the worm, what actions would you recommend to interrupt an outbreak of infection in a day care center.

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CASE STUDIES The following incidents illustrate the potential danger posed by some parasitic worms. Study the details of the cases and use your knowledge of helminth biology to answer the case study analysis questions. Trichinellosis Caused by Consumption of Wild Boar Meat—Illinois, 2013 Trichinella is a genus of tiny nematode worms that are routinely found in small mammals (such as rats) and are spread to larger predatory mammals, like pigs and bears, when the infected prey are consumed. In humans, trichinellosis—infection with Trichinella—has for many decades been linked to the consumption of pork. Changes in how pigs were raised (such as a prohibition on the use of uncooked garbage as feed stock) and the processing of pork (so as to kill the worms) led to a vast decrease in the incidence of trichinellosis in the United States over the latter half of the twentieth century. Today, most cases of trichinellosis are due to consumption of wild boar and bear meat that has been improperly prepared. On March 6, 2013, the Illinois Department of Health was notified of nine cases of trichinellosis in the Chicago area. The patients exhibited some combination of myalgia, swelling around the eyes, fever, and increased levels of eosinophils, a type of white blood cell that is generally elevated in number during parasitic infections. All nine persons had consumed meat from deer and boar that

were killed on a recent hunting trip. The meat from both animals was ground together into sausages, which were consumed by all nine persons. Trichinella spiralis larvae were identified microscopically in the sausage, but not in leftover deer meat, indicating that the boar meat was the likely source. All nine patients were treated and recovered without incident.

Anisakiasis after a Meal of Sushi—Portugal 2017 In early 2017, a previously healthy, 32-year-old man was admitted to a hospital in Portugal. Over the previous week, he had suffered severe stomach pain, vomiting, and a low-grade fever. An initial examination revealed abdominal tenderness and leukocytosis, an increase in the number of white blood cells; a collection of signs and symptoms that could point to anything from food poisoning to appendicitis. A medical history revealed that the patient had consumed sushi prior to the onset of symptoms. Endoscopy—a procedure in which a camera on the end of a long, slender tube is inserted into the gastrointestinal tract—revealed the presence of a filiform worm attached to the wall of the stomach. A Roth net, usually used to extract small batteries or other accidentally swallowed objects, was used to remove the worm, which was identified as Anisakis, a nematode. After removal of the worm, the patient’s pain immediately resolved.

CASE STUDY ANALYSIS 1. When ordering steak in a restaurant, diners are commonly asked if they would like it prepared rare, medium, or well done. When ordering pork dishes, the same question is not asked; why not?



Case Study Exercise 5  A Survey of Parasitic Worms

2. Many countries in the Middle East have very few cases of Trichinella infection. Speculate on why this could be.

3. Websites that provide tips on making your own sushi often recommend freezing the fish at very low temperatures for several hours before preparation, rather than using fresh fish. Why do you think this is?

4. What advice would you give to patients suffering from worm infections such as trichinellosis or anisakiasis to prevent spreading of the infection to roommates or family members?

REFERENCES CDC. 2014. Trichinellosis caused by consumption of wild boar meat—Illinois, 2013. Morbidity and Mortality Weekly Report, 63: 451. Carmo, J., Marques, S., Bispo, M., and Serra, D. 2017. Anisakiasis: a growing cause of abdominal pain! BMJ Case Reports 2017; doi:10.1136/bcr-2016-218857.

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NOTES

C A S E S T U DY E X E R C I S E

6

Ubiquity of Microorganisms STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Recognize microbial growth in microbiological media. 2. Differentiate between bacterial and fungal growth on solid media.

INTRODUCTION Microbiology involves, obviously, the study of microorganisms, but the history of exactly what microbes are, what they do, and where they are found has been unexpectedly complex. The century between 1850 and 1950 is often referred to as the “Golden Age of Microbiology” because this is when the importance of bacteria, protists, and fungi in our daily lives was first appreciated. For example, prior to the early 1900s, doctors, even surgeons, neglected to wash their hands between patients, their primary complaint being that it was simply too time-consuming. Saying that the “Golden Age” ended in 1950, however, was shortsighted, to say the least. Over the next half century, new bacteria continued to be discovered, viruses such as HIV and Zika became more prevalent, and entirely new infectious agents—such as the prions responsible for mad cow disease—emerged. The understanding that microbes are part of our daily life, living in

(a)

(b)

our food and drink, inhabiting our bodies, and floating in the air we breathe is not nearly as surprising now as it was a century ago. Furthermore, we realize that both the type and number of microbes are significant. We go out of our way to buy yogurt with active (bacterial) cultures while at the same time keeping milk refrigerated to prevent the growth of any more bacteria than are absolutely necessary and paying the really big bucks for mushrooms and truffles (both fungi). Microbiologists routinely use the word ubiquitous—meaning found everywhere—to describe the presence of microbes all around us. Along with specifically culturing microbes to take advantage of their abilities, we are also interested in preventing the growth of unwanted microbes, with the most obvious examples coming from the area of health care. One important difference is that in a hospital we are primarily concerned with the spread of pathogenic microbes, while in the laboratory, any unwanted microbe is capable of ruining an experiment, meaning that laboratory requirements are often more stringent than those seen in the hospital setting. The first step in determining what techniques and practices will be effective in preventing the spread of microbes is to assess their presence in the environment. Although microorganisms may be present in a variety of locations, their detection generally requires that they be removed and transferred to growth media, creating a culture, for potential identification. Today you will choose two different environmental sites to sample, using one sample to inoculate a plate of nutrient agar and the other to inoculate a tube of nutrient broth (Figure 6.1). In both

(c)

Figure 6.1  (a) A plate containing bacterial and yeast colonies. Bacterial colonies are typically smaller, shinier, and more numerous than fungal colonies. (b) Mold colonies growing on a slice of bread. Molds characteristically appear larger and more diffuse (fuzzier) than bacterial colonies. (c) Growth in a broth. The clear broth on the left is sterile while the turbidity seen in the broth on the right indicates microbial growth. No colonies will form in a liquid medium, and it is more difficult to distinguish bacterial from fungal growth. (a) ©McGraw-Hill Education/Don Rubbelke; (b) ©Steven P. Lynch; (c) ©Becton Dickinson

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cases, the idea is the same; a single bacterial cell or fungal spore, far too small to be seen on its own, is allowed to incubate on a rich growth media for several days. One cell becomes two, two become four, and so on for many generations. In a liquid medium, the growth of microbes will cause the medium, clear to begin with, to turn cloudy, or turbid. The turbidity is nothing more than the millions of cells that weren’t there a few days ago. On a solid medium within a Petri dish, the results look different because microbial cells cannot move freely through the medium as they can in a liquid. Here, a single cell produces two, then four, then eight, but the cells remain where they are, creating a mound of cells called a colony. A colony visible to the naked eye contains a million or more identical cells, all descendants of a single original cell (colonies with fewer cells certainly exist, they are just too small to be seen). The following three rules of thumb will help you discern microbial growth on a solid media: (1) You will usually have more bacterial colonies than fungal colonies, (2) bacterial colonies are usually smaller than fungal colonies, and (3) fungal colonies usually look dry and cottony, while bacterial colonies look wet and shiny, as if someone had dripped paint on the media. Also, please note the use of the word usually, three times, in that last sentence, these rules hold true most, but definitely not all, the time.

PRE-LAB QUESTIONS Remember, Understand, Apply 1. Bacterial cells and fungal spores may be found in all of the following except a. food and drink. b. indoor air. c. outdoor air. d. sterile microbiological media. 2. Microbes that are capable of causing disease are referred to as a. bacteria. b. pathogens. c. fungi. d. microbes. 3. Laboratory disinfection is generally more stringent than hospital disinfection. a. true b. false 4. When it is said that microbes are ubiquitous, it means that a. microbes in the environment are harmful. b. the immediate environment is sterile. c. only dead microbes are present. d. microbes are found virtually everywhere. 5. A tube containing liquid growth media with microbes growing within is a a. pathogen. b. culture.

c. colony. d. species. 6. A visible mound of bacteria growing on a Petri dish and containing many thousands of cells all descended from a single cell is a a. subculture. b. sample. c. colony. d. culture. 7. On a Petri dish containing both bacterial and fungal colonies, the bacterial colonies would typically be (choose all that apply) a. smaller. b. larger. c. dry in appearance. d. wet in appearance. 8. On the same Petri dish, you would generally expect _____ colonies to be greater in number. a. bacterial b. fungal 9. Bacterial growth within a liquid medium can be recognized because the medium will be a. warm. b. turbid. c. bubbling. d. clear. 10. A visible bacterial colony growing on a plate generally contains hundreds of cells. a. true b. false

PERIOD ONE MATERIALS Each student should obtain: One tube of nutrient broth One plate of nutrient agar Sterile swabs Marking pen

PROCEDURE Nutrient agar has the consistency of firm gelatin. Handle it gently to avoid breaking it. 1. For the plate of solid media, use the exposure method assigned to you in Table 6.1. For the broth tube, sample any item you like. 2. Record the appearance of the sterile media, both liquid and solid, in the results section of this exercise.



Case Study Exercise 6  Ubiquity of Microorganisms

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2. What additional information does a solid culture provide compared to a liquid culture?

TABLE 6.1 Student number

Exposure method for TSA plate

1, 10, 19

Press fingertips gently against the media

2, 11, 20

Press lips gently against media

3, 12, 21

Comb hair or scratch scalp over media for 20 sec

4, 13, 22

Blow dust onto exposed media

5, 14, 23

Expose plate to air within the laboratory for 30 min

6, 15, 24

Expose plate to air outside of the laboratory for 30 min

7, 16, 25

Expose plate to air outside the building for 30 min

8, 17, 26

Press a coin or paper money gently against the media, then remove

9, 18, 27

Any method not previously mentioned

3. Choose two objects or areas for sampling. 4. Label a tube of nutrient broth with your name, lab time, and the identity of the item or area to be sampled. 5. To inoculate liquid media, draw a sterile swab across the surface to be tested, and then transfer the swab to the media. If the surface you’ve chosen to sample is dry, dip the swab into the broth, and then use the moistened swab to sample the surface. 6. Label the bottom of a plate of nutrient agar around the edge with your name, lab time, and the identity of your second item to be sampled. 7. In the results section, record the appearance of both the broth and the agar prior to inoculation (color, clarity, consistency, etc.). 8. Incubate the inoculated media at 30°C for at least 48 h.

QUESTIONS—PERIOD ONE 1. How would you recognize microbial growth in a liquid medium? What about a solid medium?

PERIOD TWO PROCEDURE Retrieve your cultures from the incubator. Examine both the liquid media (which may require gentle shaking of the tube) and solid media. In what way have the cultures changed from their preincubation state? Exercises 41 and 42 may help you interpret any growth seen in your cultures.

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RESULTS Record your results in the table as well as on a similar table or computer spreadsheet your instructor has provided. Appearance of media prior to incubation Nutrient  broth

Nutrient  agar

Appearance of media after incubation



Case Study Exercise 6  Ubiquity of Microorganisms

Plates Plate exposure method

Colony counts Bacteria

Fungi

REVIEW QUESTIONS

Analyze, Evaluate, Create 1. What is a colony? What is the relationship between a cell and a colony?

Broth Source

Result (+/−)

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2. Based on your results, can any conclusions be drawn as to the types of habitats likely to contain the most microbes?

3. Did all the colonies on your plates look the same? Can any conclusions be drawn from this observation?

CASE STUDY The following episodes help to illustrate the growing awareness of infectious agents in the healthcare setting. Study each case carefully and use your knowledge to answer the questions which follow. Infection Control in Hospitals Florence Nightingale served as a nurse during the Crimean war in the mid-1800s, where she came to believe that one explanation for the high death rate among wounded soldiers was the spreading of respiratory secretions through long hospital corridors. She felt that open windows accelerated this process by allowing air in the wards to pass into the corridors and spread throughout the building. She also believed it was absolutely imperative that sick patients be isolated from wounded patients and worked much of her life to get these facility changes introduced in hospitals in her native Britain. Many of the observations of Florence Nightingale were incorporated into the design of Johns Hopkins Medical Center when it was built in the 1880s, a time when Ms. Nightingale’s ideas were gaining greater acceptance. The hospital was designed to isolate patients from one another, with appropriate ventilation ensuring that respiratory secretions could not easily move from room to room. One hundred and fifty years later, nearly all hospitals have come to mirror Ms. Nightingale’s vison. Air in most buildings is filtered through a HEPA (high efficiency particulate air) filter to remove fungal spores. Rooms are private or semiprivate, equipped with their own sink, and the

Long hospital corridors and large wards, like this one from the 1920s, contributed to the spread of disease by allowing microbes free passage from patient to patient. ©San Diego History Center, Historical Society



Case Study Exercise 6  Ubiquity of Microorganisms

door is fitted with a self-closing mechanism. Care is taken to ensure that surfaces (walls, ceilings, furniture, floors) are smooth and easily cleaned. Yet problems remain. From 2004 through 2006, the Toronto General Hospital reported 36 cases of infection with a multidrug resistant strain of Pseudomonas aeruginosa, with one-third of the patients dying within three months as a result of their infection. All of the patients spent time in the intensive care unit, transplant ward, or transitioning between the two. Testing of environmental specimens revealed the outbreak strain of Pseudomonas to be present in 26 sink drains spread among the three wards where infections were noted. Initial efforts to control the outbreak by sterilizing the sink, drain, and faucet yielded only temporary success, with the outbreak strain soon returning. The deep sinks and long gooseneck faucets installed in many of the hospital’s rooms were causing unexpected problems. Chosen to prevent inadvertent touching of the bottom of the bowl by hands or instruments, the sinks unfortunately were capable of contaminating other areas of the room. Using a fluorescent liquid, epidemiologists were able to show that water splashed from the sink travelled over a meter onto work surfaces and patient bedding. To reduce the risk of splashback, new faucets were installed that did not flow directly into the drain, and water pressure was decreased. A barrier was also installed between the sink and adjacent preparatory areas, while medical supplies were moved more than 1 meter from the sink. Testing with the same fluorescent liquid showed that these modifications were enough to prevent splashing water from reaching other areas, and no additional outbreak cases were seen. Between September 2014 and September 2015, three patients at the University of Pennsylvania Medical Center (UPMC) died of mucormycosis, a severe fungal infection generally restricted to people with severely compromised immune systems. The patients were organ transplant recipients and were being prescribed immunosuppressant drugs (to combat organ rejection) when they acquired their fungal infections. At the request of the United Network for Organ Sharing (UNOS) the hospital voluntarily suspended their solid organ transplantation program pending further investigation of the outbreak. The Pennsylvania Department of Health requested that the Centers for Disease Control (CDC) assist with the investigation.

Interviews were conducted with physicians, nurses, pharmacists, environmental services and facilities management, nearby construction crews, and the hospitals infection control team. Maps were created and photographs taken to highlight potential areas of risk in relation to each patient’s location at the time of diagnosis. Because construction projects commonly result in the release of fungal spores into the environment, timelines of patient hospitalization and location in comparison to the construction projects were prepared. All three patients received care in the same negative pressure isolation room within the 20-bed cardiothoracic intensive care unit (CTICU) for 14 to 58 days between their transplantation and mucormycosis diagnosis. Negative pressure rooms are usually reserved for patients with airborne infectious disease so that pathogens are retained in the room rather than being spread through the ICU. Such rooms have the potential to concentrate dust and mold spores and had already been identified as a risk factor for mold infections in immunosuppressed patients. Furthermore, the patients’ room was adjacent to a heavily used door where a carpeted hallway led to a waiting room, raising concerns that foot traffic in the area could have aerosolized mold spores that were then drawn into the negative pressure room. Staff members recalled seeing visible mold within the modular toilets present in each ICU room, and mold was also discovered behind a panel in the room, possibly related to past reports of a leaking wall-mounted dialysis fixture. As a result of their investigation, the Centers for Disease Control made several recommendations, among them: 1.  Avoid housing immunocompromised solid organ transplant patients in a negative pressure room unless otherwise indicated. 2.  Replace the frequently used door adjacent to the patients’ room with an emergency exit in order to limit foot traffic. 3.  Remove carpet from the hallway and family room adjacent to the CTICU. 4. Routinely monitor and remediate any moisture damage at the hospital. On September 27, 2015, solid organ transplants at UPMC resumed.

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CASE STUDY ANALYSIS 1. Modern hospitals often contain both semiprivate rooms and large wards where many people share a single room. In light of the spread of microorganisms, what patients would be good candidates to occupy these rooms?

2. Negative pressure rooms are sometimes referred to as source isolation rooms, while positive pressure rooms are sometime called protective isolation rooms. Explain the thinking behind these names.

3. Isolation rooms are often built with glass walls. What possible advantage does this provide?

4. Recommendations for designing new intensive care units include locating the unit near the operating theater and emergency department, but away from the main hospital wards. Why?

5. In the UPMC fungal infection outbreak, a pediatric hospital was being constructed next door to the building where the infections took place. Why is this fact important?

REFERENCES Noskin, G.A. and Peterson, L.R. 2001. Engineering Infection Control Through Facility Design. Emerging Infect. Dis. http://www.cdc.gov/ncidod/eid/vol7no2/noskin.htm Hota, S. et. al. 2009. Outbreak of Multidrug-Resistant Pseudomonas aeruginosa Colonization and Infection Secondary to Imperfect Intensive Care Unit Room Design. Infection Control and Hospital Epidemiology, 30(1):25–33. CDC. 2016. Probable Mucormycosis Among Adult Solid Organ Transplant Recipients at an Acute Care Hospital—Pennsylvania, 2014–2015. Morbidity and Mortality Weekly Report, 65:481–482.

C A S E S T U DY E X E R C I S E

7

Aseptic and Pure Culture Techniques STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Safely work with bacterial cultures. 2. Aseptically inoculate microbiological media so as to minimize the possibility of contamination. 3. Correctly isolate a pure bacterial culture using appropriate techniques.

INTRODUCTION

medical microbiologists, knew that if he wanted to prove that a specific disease was due to infection with a specific microbe, he would have to isolate the bacteria in question prior to any meaningful characterization. Techniques developed in his laboratory to produce pure cultures are still routinely used today, and mastery of these techniques is essential to success in the laboratory. Although many methods can be used to obtain a pure culture, all have the same theoretical foundation. In a successful separation, one cell in a mixed population of bacteria will be separated from all others and immobilized atop or within a solid growth medium. As this separated cell continues to reproduce over many generations, it will give rise to a single colony containing hundreds of thousands of cells, all of which are derived from a single progenitor (Figure 7.1). Each colony can now be used for further study of the bacterium. In this exercise, three different techniques are provided to accomplish the isolation; your instructor will tell you which procedures you will perform.

Studying microorganisms is fundamentally different than studying any other type of living organism. Microorganisms are found virtually everywhere, including soil, water, air, dust, food, and most body surfaces, even when the body is healthy. Furthermore, these same microbes are generally invisible to the naked eye, meaning their presence is rarely apparent, even when a sample is heavily popMixture of cells in sample ulated with microbes. The combination of the ubiquitous nature of microbes and their “invisibility” means that the microbiologist must use specific aseptic techniques to exclude unwanted Separation of microbes. The proper use of these techniques Microscopic view cells by spreading Parent cannot be overemphasized; without them the or dilution on agar cells study of microbiology is a hopeless exercise. medium Additionally, these techniques must be mastered Incubation quickly: the organisms you work with will not care that you’ve only recently begun to study Growth increases the number of cells microbiology! The proper use of aseptic techniques to reduce the occurrence of contamination in the laboratory is only part of the story, however. It Microbes become is an unfortunate fact of life that bacteria must visible as isolated Macroscopic view be studied as a single species but are rarely colonies containing millions of cells found this way in the environment. In the real world, whether it is the normal microbiota of the body, a wound, sewage, soil, or water, bacteria are found in mixed populations made up of many different species. Properly studying a sin- Figure 7.1  Regardless of the method used, all pure culture techniques depend on gle species however, requires that it be isolated separating individual cells of a culture from one another. Once separated, each cell from other species in the population to obtain a will produce a colony containing many hundreds of thousands of cells, all of which are pure culture. Robert Koch, one of the earliest descendents of the single isolated cell.

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Case Study Exercise 7  Aseptic and Pure Culture Techniques

PRE-LAB QUESTIONS Remember, Understand, Apply 1. Microorganisms are commonly found in soil, surfaces, and dust, but not on living surfaces like skin. a. true b. false 2. Microbes are generally (choose all that apply) a. found everywhere. b. invisible to the naked eye. c. only present when a person is ill. d. not found within the air of a laboratory. 3. When a sample is heavily populated with microbes, the microbes are generally visible to the naked eye. a. true b. false 4. Laboratory techniques that exclude unwanted microbes are called _____. a. ubiquitous b. aseptic c. invisible d. manipulations 5. Isolation techniques are necessary in the microbiology lab because a. microbes grow better by themselves. b. microbes are usually found as part of a mixed population. c. microbes are not infectious individually. d. microbes will infect one another if they are not separated. 6. A tube of nutrient agar, upon which is growing a single species of bacteria is said to be a _____ culture. a. pure b. mixed c. contaminated d. pathogenic 7. A mound of hundreds of thousands of bacterial cells growing on a plate of agar, all descended from the same original cell is a a. mixture. b. spore. c. pathogen. d. colony.

PROCEDURE

General Considerations The following practices should be followed under all circumstances. They are intended to promote safety in the laboratory and reduce contamination. ∙∙ Keep your work area uncluttered and organized. Your benchtop will most likely be crowded with inoculating tools, stains, test tube rack, Bunsen burner, lab book, notebook, etc. Keep other items to a minimum. Label tubes as instructed; this may include using tape, paper held on with rubber bands, or laboratory marker. Petri dishes may be labeled directly on the base of the plate. ∙∙ Test tubes should always be placed in a test tube rack when not in use. Use the rack to transport the tubes as well. Plates should be carried in a sleeve if there are more than can be comfortably carried by hand, generally two or three. (Figure 7.2) ∙∙ Regulate the flame of your Bunsen burner by altering the amount of air entering the bottom of the burner. Adjust the amount of air entering the burner until the flame has both an inner and outer cone. The hottest part of the flame is the tip of the inner blue cone, and this is the area that is used for sterilization of inoculating tools. The outer, less intense

PART I: ASEPTIC TECHNIQUE There are no hands-on exercises for this portion of the lab, but these procedures should be used whenever you are working with microorganisms.

Figure 7.2  Metal sleeves, or similar containers, should be used whenever you have more than a few Petri dishes. Tubes of media should always be carried in racks to ensure that they remain upright. ©Barry Chess



Case Study Exercise 7  Aseptic and Pure Culture Techniques

Figure 7.3  The hottest portion of a Bunsen burner’s flame is found at the edge of the inner blue cone. Inoculating loops and needles should be sterilized in this area. The area outside the cone is slightly cooler but is acceptable for flaming of tubes during inoculations. ©Barry Chess

∙∙ ∙∙

∙∙

∙∙

∙∙

flame is adequate for flaming of tubes and heat fixing of bacterial smears (Figure 7.3). Inoculating loops and needles should be held lightly with your dominant hand, while culture tubes should be held with the nondominant hand. Sterilize inoculating tools by passing them through the flame until each portion of the wire has achieved an orange color. Begin flaming about 2–3 cm up the handle and draw the tool backward through the flame until each part of the wire has been heated to a uniform orange. Flaming from handle to tip allows the end of the wire to heat up slowly and helps prevent the formation of bacteria-containing aerosols. Remove caps from tubes by grasping them with the smallest two fingers of your dominant hand and using these fingers to hold the cap while inoculating. Tight fitting or screw-on caps may have to be loosened before beginning this procedure (Figure 7.4). The neck of an open tube should be sterilized immediately after removing the cap and once again prior to replacing it. This can be accomplished by passing the tube briefly through the flame of the burner. Open tubes should always be held at an angle to minimize the possibility of airborne contamination (see Figure 7.4). Prior to transfer, inoculating loops and needles must be allowed to cool for 10–15 sec. If a hot inoculating tool is

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Figure 7.4  The last two fingers of your dominant hand should be used to hold the cap of any tube you are working with. The remaining three fingers are used to manipulate the inoculating tool. ©Barry Chess

Figure 7.5  When working with a Petri dish, the lid should be used as a shield to prevent contamination of the medium by unwanted organisms. ©Barry Chess

plunged into media containing bacteria, an aerosol may be created, releasing bacteria into the air. ∙∙ When transferring organisms to or from a tube, hold the inoculating tool steady and move the tube. Reducing the movement of the loop or needle lessens the possibility of bacteria ending up where they are not wanted. ∙∙ When transferring organisms to or from a Petri dish, use the lid to protect against airborne contamination (Figure 7.5).

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Case Study Exercise 7  Aseptic and Pure Culture Techniques

Figure 7.7  A needle is used to remove a small quantity of bacterial growth from a slant. ©Barry Chess

Figure 7.6  Gently mix broth cultures in order to suspend the organisms within. On the left, the bacteria are found in a mass on the bottom of the tube; while on the right, the bacteria are well distributed throughout the media. ©Barry Chess

3. Holding the needle still, move the tube up and around the needle until it is positioned above the growth. Carefully touch the needle to the bacteria, obtaining a tiny (often invisible) quantity of growth. Withdraw the tube from the needle (Figure 7.7). 4. Flame the neck of the tube, and replace the cap. 5. Transfer the bacteria to the intended medium.

Obtaining Growth from an Agar Plate

Specific Transfer Methods Because the same aseptic techniques are applicable in virtually all transfers, specific procedures will vary little from situation to situation. The most important aspect is the choice of inoculating tool, which allows the appropriate amount of bacteria to be transferred. Typically, a loop is used to remove bacterial growth from a liquid culture and a needle is used to remove growth from a solid culture, such as a slant or a plate. Notice that the medium to which growth will be transferred is not usually a concern. Although abridged step-by-step instructions follow, be sure to use the proper aseptic technique as previously outlined.

Obtaining Growth from a Broth 1. Suspend bacterial growth in a broth tube by tapping the tube with your fingers until a vortex of cells is seen swirling in the tube. A properly mixed culture will be turbid (Figure 7.6). 2. Flame an inoculating loop, and allow it to cool. 3. Remove the cap from the broth tube, and flame the neck of the tube. 4. Holding the loop still, move the tube up and around the loop until the loop is submerged in the broth. Carefully withdraw the tube from the loop. 5. Flame the neck of the tube, and replace the cap. 6. Transfer the bacteria to the intended medium.

Obtaining Growth from a Slant 1. Flame an inoculating needle, and allow it to cool. 2. Remove the cap from the slant, and flame the neck of the tube.

1. Flame an inoculating needle, and allow it to cool. Touching the needle briefly to an uninoculated portion of the plate will ensure that it is cool. 2. Remove the cover from an agar plate, but hold it over the plate to act as a shield against airborne contamination (see Figure 7.5). 3. Carefully touch the needle to an isolated bacterial colony, obtaining a tiny (often invisible) quantity of growth. Withdraw the needle from the plate, and replace the cover. 4. Transfer the bacteria to the intended media.

Inoculating an Agar Slant with a Fishtail Streak 1. Remove the cap from the slant, holding it with the smallest finger of your dominant hand. 2. Flame the neck of the tube. 3. Keeping the tube tilted (to guard against airborne contamination), slide the tube up and around the needle or loop. The surface of the agar inside the tube should be facing upward. 4. Touch the needle or loop to the surface of the agar, and while moving the tool in a zigzag pattern, slowly withdraw the tube (Figure 7.8). 5. Flame the neck of the tube, and replace the cap. 6. Sterilize the inoculating tool in the flame of the Bunsen burner.

Inoculating an Agar Deep 1. Remove the cap from the deep, holding it with the smallest finger of your dominant hand. 2. Flame the neck of the tube.



Case Study Exercise 7  Aseptic and Pure Culture Techniques

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Figure 7.8  A loop is used to inoculate an agar slant with a broth culture. ©Barry Chess

3. Keeping the tube tilted (to guard against contamination), slide the tube up and around the needle. 4. Insert the needle into the agar until it is about 1 cm from the bottom of the tube (Figure 7.9). Carefully remove the needle from the agar so that it follows the same pathway exiting the agar as it did entering it. 5. Flame the neck of the tube, and replace the cap. 6. Sterilize the inoculating needle in the flame of the Bunsen burner.

Inoculating a Broth 1. Remove the cap from the broth tube, holding it with the smallest finger of your dominant hand. 2. Flame the neck of the tube. 3. Keeping the tube tilted (to guard against airborne contamination), slide the tube up and around the needle or loop until it is submerged in the broth. Swirl the inoculating tool several times to ensure the transfer of the cells. 4. Slide the tube away from the inoculating tool. If using a loop, touch it to the side of the tube to remove as much broth as possible (Figure 7.10). This reduces the chance of aerosol formation when the loop is flamed and the

Figure 7.10  When using a loop to sample a liquid culture, touching the loop to the inner surface of the tube momentarily will remove excess broth, reducing the chances of a dripping culture contaminating your work surface and the formation of aerosols when the loop is flamed. ©Barry Chess chances of a dripping culture contaminating your work surface. 5. Flame the neck of the tube, and replace the cap. 6. Sterilize the inoculating tool in the flame of the Bunsen burner.

PART II: PURE CULTURE TECHNIQUES

PERIOD ONE Three methods are described here: the streak-plate method, the loop dilution method, and the spread-plate method. Your instructor will tell you which method(s) to use.

I. Streak-Plate Method This method is the most economical in terms of time and materials, requiring just a few minutes and only a single plate of media. Its main drawback is that a certain degree of skill is required, which takes time to fully develop. The streak-plate method of isolation is outlined in Figure 7.11.

MATERIALS

Figure 7.9  A needle is inserted nearly to the bottom of the medium when inoculating a deep. Note that the tube is held nearly parallel to the work surface, which reduces the chance of contamination. ©Barry Chess

Each student should obtain: A mixed broth culture containing: Escherichia coli Serratia marcescens Micrococcus luteus One nutrient agar plate Inoculating loop Marking pen

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Case Study Exercise 7  Aseptic and Pure Culture Techniques

1st

1st

(a) 1st

2nd

(a)

(b)

(c)

I

II

III

I

II

III

(b) 2nd

1st

2nd (d) 3rd

3rd

4th (c)

(d)

Figure 7.11  (a) In a streak plate, a single loopful of a culture is spread four or five times in the first quadrant. (b) The loop is flamed and allowed to cool for about 10 sec. Touching the loop to an uninoculated area of the plate will ensure that it is cool. Once cool, five or six streaks are made from quadrant 1 into quadrant 2. (c) Following flaming and cooling of the loop, six to seven streaks are made from quadrant 2 into quadrant 3. (d) After flaming and cooling once more, several streaks are made from quadrant 3, using up all of the uninoculated space on the plate. Finally, the loop is flamed before being placed aside.

PROCEDURE 1. Label the bottom of your Petri dish with your name and lab time. Place the label around the periphery of the plate so your view of the bacterial colonies will be as complete as possible after the incubation. 2. Carefully agitate the tube containing the mixed culture until the bacteria are suspended in the medium. 3. Flame the loop to red hot, and allow it to cool. Remove the cap from the culture tube, and while holding the cap with the pinky finger of your dominant hand, flame the neck of the tube. 4. Remove a single loopful of broth from the tube. 5. Flame the neck of the tube and replace the cap. 6. Streak the organism onto the plate as shown in Figure 7.11. Hold the Petri dish cover over the plate to guard against airborne contamination as you work. Use as little pressure as possible to avoid gouging the medium. Begin by applying 4 or 5 parallel streaks in the first quadrant. 7. Flame the loop, and allow it to cool. Touching the loop briefly to an uninoculated area of the plate will ensure that it has cooled sufficiently. Make 5 or 6 streaks extending from the first quadrant to the second. Flame the loop, and allow it to cool. 8. Make 6 or 7 streaks extending from the second quadrant to the third. Flame the loop, and allow it to cool.

Figure 7.12  (a) In a loop dilution, a single loopful of a culture is used to inoculate a tube of melted agar that has been cooled to 50°C. The loop is used to stir the medium so that bacterial cells are spread evenly throughout the tube. (b) After flaming the loop and allowing it to cool for about 10 sec, a loopful of medium from tube I is removed and used to inoculate tube II. Again, the loop should be used to distribute the bacteria throughout the tube. (c) The loop is flamed and allowed to cool once again, and then used to transfer a loopful of medium from tube II to tube III. Flame the loop before setting it down. (d) Once all three tubes have been inoculated, flame the neck of tube I and pour the agar into the bottom of the first Petri dish, carefully swirling the dish to ensure that the medium covers the entire bottom of the plate. Do the same for tubes II and III. 9. Make as many streaks as is practical, beginning in the third quadrant and extending outward. Use as much of the uninoculated area of the plate as possible but do not allow the loop to enter either of the first two quadrants. 10. Flame the loop before placing it aside. 11. Incubate the plate in an inverted position at 30° C for 72–96 h. Inverting the plates prevents condensation (which may have accumulated on the lid of the plate) from dropping onto the agar and causing the organisms to spread over the agar surface, ruining the entire isolation procedure.

II. Loop Dilution (Pour Plate) Method This method consumes more time and materials than does a streak plate, but the results produced are generally quite good. The loop dilution method of isolation is outlined in Figure 7.12.

MATERIALS Each student should obtain: A mixed broth culture containing: Escherichia coli Serratia marcescens Micrococcus luteus Three nutrient agar pours Three empty Petri dishes Hot plate Inoculating loop Thermometer Marking pen



Case Study Exercise 7  Aseptic and Pure Culture Techniques

PROCEDURE 1. Label the bottom of your Petri dishes with your name and lab time. Also label each dish I, II, or III. Place the label around the periphery so your view of the plate will be as complete as possible after the incubation. 2. Label the agar tubes I, II, and III, and place them into a beaker containing enough water to cover the agar in the tubes. Bring the water in the beaker to a boil, and allow 5 min for the agar to liquefy. Remove the beaker from the hot plate, place a thermometer in the water, and allow the agar to cool to 50°C. Adding fresh water to the beaker will allow it to cool faster. Leave the melted agar tubes in the warm water as this will maintain them at 50°C. 3. Carefully agitate the tube containing the mixed culture until the bacteria are suspended in the medium. 4. Flame the loop to red hot, and allow it to cool. Remove the cap from the culture tube, and while holding the cap with the pinky finger of your dominant hand, flame the neck of the tube. 5. Remove a single loopful of broth from the tube. 6. Flame the neck of the tube, and replace the cap. 7. Remove the cap from tube I, and flame the neck of the tube. Inoculate the medium by submerging the loop into the agar and gently swirling it to dislodge and mix the bacterial cells. After removing the loop, flame the neck of the tube and replace the cap. Place the tube back in the water-filled beaker. 8. Flame the loop and allow it to cool for 5–10 sec. Remove the cap from tube I, and flame the neck of the tube. Carefully remove a single loopful of inoculated agar. Flame the tube, replace the cap, and return the tube to the beaker. 9. Remove the cap from tube II, and flame the neck of the tube. Inoculate the medium by submerging the loop into the agar and swirling it to dislodge and mix the bacterial cells. After removing the loop, flame the neck of the tube and replace the cap. Return the tube to the water-filled beaker. 10. Flame the loop and allow it to cool for 5–10 sec. Remove the cap from tube II, and flame the neck of the tube. Carefully remove a single loopful of inoculated agar. Flame the tube, replace the cap, and return the tube to the beaker. 11. Remove the cap from tube III, and flame the neck of the tube. Inoculate the medium by submerging the loop into the agar and swirling it to dislodge and mix the bacterial cells. After removing the loop, flame the neck of the tube and replace the cap. Return the tube to the water-filled beaker. Flame the loop before placing it aside. 12. Remove the cap from tube I, and flame the neck of the tube. Carefully pour the liquefied agar into the bottom of plate number I. Gently swirl the plate until the agar has completely covered the base of the plate. Flame the neck of the tube, and replace the cap. Repeat this process for tubes II and III. 13. After the media have completely solidified, incubate the plates in an inverted position at 30°C for 72–96 h.

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III. Spread-Plate Method This method gives consistently reliable results when bacterial samples are dilute or if the medium being inoculated is highly selective and allows only a limited number of cells to grow. If a culture has an abundance of bacterial growth, it should be diluted with sterile water prior to spreading, or a different technique should be used. The spread-plate method of isolation is outlined in Figure 7.13.

MATERIALS Each student should obtain: A previously diluted mixed broth culture containing: Escherichia coli Serratia marcescens Micrococcus luteus One nutrient agar plate Approximately 50 ml of ethyl or isopropyl alcohol in a 250-ml beaker A large beaker that will completely cover the small beaker and spreading rod Bent glass spreading rod One sterile dropper or pipette Sterile water Marking pen Petri plate turntable (optional)

PROCEDURE A Safety Note: Should the alcohol in the beaker catch fire, place the larger beaker over the alcohol-containing beaker and spreading rod. When the oxygen is exhausted, the fire will extinguish itself. 1. Place the spreader rod in the beaker, ensuring that the alcohol covers the lower portion of the rod. 2. Arrange the items on your bench so that the Bunsen burner is between the alcohol and the Petri dish. This reduces the chance of accidentally setting the alcohol aflame. 3. Mark the bottom of your Petri dish with your name and lab time. Place the label around the periphery so your view of the plate will be as complete as possible after the incubation. 4. Carefully agitate the tube containing the diluted mixed culture until the bacteria are suspended in the medium. 5. Place a single drop of sterile water in the center of the plate. Flame your loop, and retrieve a single loopful of the dilute culture. Add this to the drop of water on the plate. Flame the loop prior to placing it aside. 6. Holding the upper end of the spreader rod, remove it from the alcohol and pass it through the flame of the Bunsen burner, allowing the alcohol to ignite. Be sure to keep your hand above the spreader so that the flaming alcohol doesn’t run onto your hand. Wait until the alcohol has completely burned off before continuing.

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Case Study Exercise 7  Aseptic and Pure Culture Techniques

(a)

7. Place the lower portion of the spreader flat against the agar plate. Rotate the plate using your thumb and middle finger while moving the spreader back and forth. If a turntable is used, gently spin the turntable while moving the spreader back and forth. In both cases, finish by rotating the plate one complete revolution while holding the spreader against the edge of the plate. 8. Return the spreader to the alcohol. There is no need to flame it again. 9. Allow the plate to sit upright for 5 min, then incubate it in an inverted position at 30°C for 72–96 h.

PERIOD TWO—Results (b)

(c)

Retrieve your plates from the incubator. To make differentiation easier, each bacterium used in this exercise exhibits a unique color. Escherichia coli produces off-white growth, Micrococcus luteus produces yellow growth, and Serratia marcescens produces red growth. Rarely, if ever, will you encounter a mixed culture with such vibrant colors outside the laboratory, but here the colors will help you to more fully evaluate the success of your separation. Regardless of the method used, a successful outcome will always be marked by many well-isolated colonies (­Figure 7.14).

(d)

(a)

(e)

Figure 7.13  (a) When preparing to isolate bacteria using a spread plate, arrange your bench so that the Bunsen burner is between the alcohol and the Petri dish. This minimizes the chances of setting fire to the alcohol. (b) Use a Pasteur pipette to add a single drop of sterile water to the surface of the media. (c) Flame your loop and allow it to cool for about 10 sec, then use it to inoculate the water drop with a loopful of the mixed culture. (d) Remove the glass spreading rod from the alcohol and pass it through the flame, allowing the alcohol to ignite. Remove the rod from the flame and wait for the alcohol to burn off completely. (e) Place the plate on a turntable and rotate the turntable while sliding the spreader back and forth across the surface of the medium. If a turntable is not available, the plate can simply be rotated on the benchtop. In either case, finish the inoculation by rotating the plate a final turn while holding the spreader against the edge of the plate. After inoculating the plate, return the spreader to the alcohol; there is no need to flame it again.

(b)



Case Study Exercise 7  Aseptic and Pure Culture Techniques

89

Loop dilution:

Spread plate:

(c)

Figure 7.14  (a) Completed streak plate displaying well-isolated colonies of Serratia marcescens (red), Escherichia coli (white), and Micrococcus luteus (yellow). (b) Loop dilution of the same three bacterial species. Elongated (football shaped) colonies are those that have developed within the agar rather than on the surface of the medium. (c) Spread plate of E. coli and M. luteus. (a, c) ©Kathy Park Talaro; (b) ©Barry Chess

If your plate looks beautiful, all the better, but as long as several well-isolated colonies are present, the plate can be judged a success. Record the appearance of your plates for each separation technique you employed. Critically analyze your plate(s), rating each on a scale of 1 (unusable) to 10 (­Microbiology Hall of Fame). For anything less than a 10, indicate how the plate could have been improved. Streak plate:

Subculturing Bacterial Isolates Although isolated colonies may be used directly from the Petri dishes on which they were isolated, in most cases, it is advisable to transfer, or subculture, an isolated colony to its own container of media. This allows for easier study and organization of bacterial isolates. Today you will subculture each of your isolated bacterial species to nutrient agar slants. This procedure is illustrated in Figure 7.15.

MATERIALS Each student should obtain: Bacterial plates containing isolated colonies Three nutrient agar slants Bunsen burner Inoculating needle Marking pen

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Case Study Exercise 7  Aseptic and Pure Culture Techniques

PROCEDURE

(a)

1. Using any combination of plates, identify one well-isolated colony for each bacterial species. On the bottom of the Petri plate, circle the colony so as to make it easier to find when you are ready to inoculate. 2. Label each agar slant with its respective bacterium, E. coli, S. marcescens, or M. luteus. 3. Flame the inoculating needle completely, including 2–3 cm of the handle, and allow it to cool. 4. Open the Petri dish containing a well-isolated E. coli colony, and touch the tip of the needle to the colony you selected earlier. Replace the cover of the Petri dish. 5. Remove the cap from the agar tube labeled E. coli and flame the neck of the tube. 6. Inoculate the slant using a fishtail streak. Flame the neck of the tube, and replace the cap. Flame the inoculating needle prior to setting it aside. 7. Repeat steps 4 through 6 for the other two bacteria. 8. Incubate the slants at 30°C for 48 h.

PERIOD THREE Retrieve your slants from the incubator, and record their appearance, rating them as you did your plates. (b)

(c)

Figure 7.15  (a) For each species of bacteria, identify a single wellisolated colony. (b) Flame your inoculating needle and allow it to cool. Touch the needle to the isolated colony, picking up a minute amount of bacterial growth. (c) Flame the neck of the nutrient agar tube and inoculate the tube by drawing the needle along the surface of the agar, from bottom to top, moving the needle back and forth as it ascends up the surface of the slant. Flame and recap the tube, and flame the needle before setting it aside. Repeat this process for each of the other two species.



Case Study Exercise 7  Aseptic and Pure Culture Techniques

REVIEW QUESTIONS Analyze, Evaluate, Create 1. What is the importance of generating isolated bacterial colonies?

2. Describe how a bacterial sample would be obtained from and inoculated into each of the following types of media. Agar slant:

Agar plate:

Broth:

3. Draw the results you would expect to see in a well-done streak plate, loop dilution, and spread plate.

streak plate

4. What is a subculture?

loop dilution

spread plate

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Case Study Exercise 7  Aseptic and Pure Culture Techniques

5. Sketch the results you would expect to see for three well-isolated subcultures.

6. How would a subculture appear if a colony containing both S. marcescens and M. luteus was subcultured to a slant?

7. Condensation often gathers in the bottom of agar slants. Why is it important in this exercise to limit condensation on plates but not on slants?

8. Which separation method is not appropriate for use with cultures containing a great deal of bacterial growth? Why is this method not a good choice for these conditions?

9. Why is agar cooled to 50°C prior to being inoculated with bacteria? What would happen if the agar were significantly warmer or cooler when inoculated?

10. How could you identify a potential contaminant on a streak plate? A pour plate? A spread plate?

11. How would any of the isolation techniques seen in this laboratory be affected by the use of a selective medium?



Case Study Exercise 7  Aseptic and Pure Culture Techniques

93

CASE STUDY In microbiology, the need to obtain a pure culture is simply a fact. When outbreaks occur, the single microbe responsible must be isolated, and in the world outside the laboratory, this is often easier said than done. The following case emphasizes the necessity of obtaining a pure culture and the twists and turns that an outbreak investigation may take. Study the case and answer the questions that follow.

Generating Questionnaire, a lengthy survey with questions about past travel, re­cent meals, hospital stays, and animal contact. Examination of credit card receipts, frequent-buyer cards, and social media interactions revealed that the two pa­ tients had fresh produce purchases from the same grocery store and had six common restaurant exposures. On October 19, CDPHE was notified of a third person who had tested positive for Typhoid Fever—Colorado, Salmonella Typhi infection, 2015 with symptoms beginning Fresh food, because it is never subjected to the protective heat that Typhoid fever is caused by in- cooking provides, has the potential to harbor pathogenic bacteria. September 15. An investifection with Salmonella en- ©CampPhoto/iStock/Getty Images gation similar to that used terica, serovar Typhi—often for the first two cases was shortened to Salmonella Typhi—the most worrisome strain undertaken, and it revealed that all three patients had eaten of Salmonella. Rare in the United States, Salmonella Typhi at the same location, a Mexican restaurant where food was is endemic in many countries, where it is responsible for prepared from fresh ingredients, the only epidemiological 22 million infections and 200,000 deaths per year. Of the link between all three cases. 5700 cases of typhoid fever reported yearly in the United Inspections performed within the restaurant revealed no States, nearly all are seen in people who have recently travdeficiencies with regard to hand hygiene or food handling. elled internationally. Humans are the only reservoir for SalmoRestaurant administrators cooperated with the investigation, nella Typhi; and the bacterium is typically transmitted when supplying the names of all food handlers, who were given feces from an infected person comes into contact with food paid time off to be interviewed and examined. Because no or water. employees were symptomatic, investigators presumed they On September 11, 2015, a single case of typhoid fever were searching for a carrier, someone infected with a pathowas reported to the Colorado Department of Public Health gen who doesn’t display signs or symptoms of disease. The and Environment (CDPHE). Because this patient had recently normal microbiome of the human gut contains many hunreturned from a trip abroad, the infection was at first thought dreds of different species of bacteria, so isolating one particto be travel-associated, but when a second case was identiular species can be a daunting task. Samples were obtained fied, in a patient with no recent travel, the CDPHE initiated an from the rectum of each employee, and pure culture techinvestigation with three goals: (1) to determine whether these niques were used to isolate and identify Salmonella Typhi in cases represented part of a larger outbreak, (2) to identify a single food handler. This employee reported having travcommon exposure sources, and (3) to stop transmission of elled 15 years previously to a country where typhoid fever the bacterium. was endemic, but he had not been ill, and he had not had The patients first displayed symptoms—fever, headache, contact with any ill persons. The worker was excluded from constipation, chills, myalgia, and malaise—on September 2 food service work and treated with azithromycin for 28 days. and 20, respectively. Initial questioning revealed no immeAfter three consecutive stool specimens (obtained at least one month apart) tested negative for Salmonella Typhi, the diate epidemiological link; the patients did not know one employee was allowed to return to work, where his job had another, and they lived 6 miles apart. Investigators quesbeen held open for him. tioned each patient using the Salmonella National Hypothesis

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Case Study Exercise 7  Aseptic and Pure Culture Techniques

CASE STUDY ANALYSIS 1. Why was it important in this case to identify Salmonella Typhi in the feces of the restaurant worker? Wouldn’t the discovery of any bacterium be adequate?

2. Which of the three isolation techniques in this exercise would have been least suited to isolation of Salmonella Typhi in this case? Why?

3. MacConkey agar is a selective medium that only allows certain types of bacteria, including Salmonella, to grow. How could the use of MacConkey agar have simplified the isolation of Salmonella Typhi in this case?

4. Thinking through the case, other than the restaurant workers, when else was microbiological sampling likely used?

REFERENCE CDC. 2016. Typhoid Fever Outbreak Associated with an Asymptomatic Carrier at a Restaurant—Weld County, Colorado, 2015. Morbidity and Mortality Weekly Report, 65:606–607.

C A S E S T U DY E X E R C I S E

8

Simple Staining, Negative Staining, and Gram Staining Diplococcus (2 cells) Streptococcus (variable number of cells)

STUDENT LEARNING OUTCOMES

1 plane (a)

After completing this exercise, you should be able to: 1. Prepare a bacterial smear from both solid and liquid cultures. 2. Properly stain bacterial cultures using simple-, negative-, and Gram-staining techniques.

INTRODUCTION At its most basic, microbiology deals with the differences betweens cells. Microscopic examination is used to determine a cell’s size and shape as well as the often characteristic arrangement the cells of a given species assume as they multiply. The initial classification of an isolate is absolutely dependent upon the cells’ shape, arrangement, and staining characteristics, although complete identification is not possible from these aspects alone. Cells of a given species all generally have the same shape, or morphology. Bacterial cells are classified as cocci (singular, coccus) if they are spherical in shape, rods or bacilli (singular, bacillus) if they are elongated, and ­spirilla (singular, spirillum) if they are spiral shaped. Although division based on shape holds true in most instances, variations on this theme do exist. Short curved rods (­vibrios), cells intermediate between rods and cocci (coccobacilli), and flexible spiral bacteria (spirochetes) are all seen. P ­ leomorphic bacteria exhibit a variety of shapes, even in the same sample. Cellular arrangement, the manner in which bacterial cells associate with one another, is a by-product of the manner in which a cell divides and whether the cells stay attached after division. Cocci produce more arrangements than rods. If a coccus divides along a single plane and the cells remain attached after division, a diplococcus is formed. If the cells continue to divide in this way without separating, a streptococcal arrangement results. If, instead, a second division takes place perpendicularly to the first, a four-cell group called a tetrad is produced while a third division (perpendicular to the first two) results in a cuboidal packet of eight cells called a sarcina. If the planes of division are not at right angles to one another, an irregular cluster of cells called a staphylococcal arrangement results (Figure 8.1). Rod-shaped bacteria divide only along the transverse plane (i.e., the shortest possible distance across the cell),

Tetrad (4 cells) Sarcina (packet of 8–64 cells) or

2 perpendicular planes (b) Irregular clusters (number of cells varies) Several planes (c)

Staphylococcus and Micrococcus

Figure 8.1  Arrangement of cocci resulting from different planes of cell division. (a) Division in one plane produces diplococci and streptococci. (b) Division in two planes at right angles produces tetrads and packets. (c) Division in several planes produces irregular clusters.

producing the only arrangements seen in the bacilli, diplobacilli, streptobacilli, and palisades (Figure 8.2). Spiral-shaped bacteria rarely remain attached after division and consequently are seen only as single cells. In most cases, a sample will have multiple arrangements visible, for instance, diplococci, staphylococci, and tetrads. In these cases, the most prevalent arrangement tends to be emphasized because it usually represents the end result of the growing bacterial grouping. Bacterial cells are essentially transparent, and a careful examination of them involves the same problems as a careful examination of ice cubes in a glass of water. Because the cells and their background do not contrast well, they are difficult to visualize. Colored stains are used to increase the contrast between bacterial cells and their background, and while these stains differ in many ways, they share a basic chemistry that allows them to interact with cells. The colored portion of a stain molecule is known as the chromophore, and this portion of the molecule also typically carries an electrical charge. Ionic interactions between chromophores and cells are responsible for coloring the cell. Staining techniques in the microbiology laboratory have been designed to increase the contrast between bacterial cells and their background in one of two ways. In negative ­staining, the background is stained, leaving clear cells visible against a dark background while in simple staining, the cells are stained, leaving them visible as dark objects against a light background. Both types of staining are seen in Figure 8.3. Negative staining uses negatively charged dyes such as nigrosin or

95

96

Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

(a)

(a)

(b)

(b)

Figure 8.3  (a) In negative staining, the background is stained but the cells are not, resulting in light organisms against a dark field (1000x). (b) In simple staining, bacterial cells are stained directly and appear as dark colored cells against a light background (1000x). (a) ©McGraw-Hill Education/Lisa Burgess, photographer; (b) ©Lee W. Wilcox

(c)

Palisades arrangement

Snapping

Figure 8.2  Arrangements of rod-shaped bacteria. (a) Bacillus anthracis displays a chain of cells (1000×). (b) “Snapping” of Corynebacterium diptheriae (28,000×) cells leads to (c) a palisades arrangement (1200×). (a) Source: CDC; (b) ©Dr. Gary Gaugler/Science Source; (c) Source: CDC

India ink, both of which are repelled by the negative charge of the bacterial cell but are attracted to the glass slide. In simple staining, any positively charged, or basic, stain is used to stain the negatively charged bacterial cell. Common basic stains in the microbiology laboratory include methylene blue, crystal violet, malachite green, and safranin (which is pink). Although simple and negative staining increase the contrast between a cell and its background, making it easier to determine the size,

shape, and arrangement of a particular bacterium, they provide little information beyond this. One advantage seen with negative staining is a more accurate determination of the size of a bacterial cell. When preparing a cell for simple staining, heat is applied to the bacterial sample, both to kill the cells and to affix them to the slide. An unwanted consequence of heating is that cells generally shrink. Because negative staining uses no heat, cell shrinkage is minimized or eliminated. A differential stain, as the name implies, allows the discrimination of one cell from another based on differential-staining properties. These stains provide not just morphological information, such as size, shape, and arrangement, but also give a clue as to the biology of the cell by highlighting structural differences that exist between groups of bacteria. Although several types of differential stains exist, the most widely used, and in fact the most important stain in all of microbiology, is the Gram stain, developed in 1884 by the physician Hans Christian Gram. This technique separates bacteria into two groups, Gram positive and Gram negative, based on differences in the structure of the cell wall. The steps in a Gram stain are outlined



Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

Reagent

Gram positive

Gram negative

None (Heat-fixed cells) Crystal violet (30 sec) Gram’s iodine (1 min) Ethyl alcohol (10–20 sec) Safranin (1 min)

Figure 8.4  Color of cells after each step of the Gram-staining

97

∙∙ Cells are decolorized with ethyl alcohol. The alcohol dissolves the lipids that are found in the outer membrane of Gramnegative cells, allowing the crystal violet–iodine complex to escape. At this point, Gram-negative cells are colorless while Gram-positive cells are still dark purple. ∙∙ Safranin, a counterstain, is then added to increase the contrast of the colorless Gram-negative cells, rendering them pink. The Gram-positive cells are dyed pink as well, but the darker color of the crystal violet masks the lighter color of the safranin. In the end, those cells staining purple are designated Gram-positive while those staining pink are deemed Gramnegative. The ability to reliably Gram stain is absolutely essential to the correct identification of a bacterial species, and reliability comes about only with much practice. A variety of bacteria, stained using the techniques presented in this exercise, may be seen in the bacterial gallery, Figure 8.5.

procedure.

here, and the appearance of the cells at each point in the process is illustrated in Figure 8.4: ∙∙ Cells are stained with the primary stain crystal violet. Both Gram-positive and Gram-negative cells are colored a deep purple at this point. ∙∙ Iodine is added as a mordant, a chemical that serves to fix a dye in a staining process. In this case, the iodine binds with the crystal violet to create an insoluble complex within the thick peptidoglycan layer of Grampositive cells.

(b) Gram stain of Bacillus cereus subspecies mycoides (1000x). This species of Bacillus commonly causes food poisoning. Although classified as Gram-positive, many species of Bacillus will begin to stain Gram-negative, as seen here, when the culture ages.

(a) Gram stain of Bacillus anthracis (1000x). The causative agent of anthrax in humans and animals, B. anthracis often forms long chains in culture.

Figure 8.5  (a) Source: CDC; (b) Source: CDC/Dr. William A. Clark; (c) ©McGraw-

Hill Education/Lisa Burgess, photographer; (d) Source: CDC/Dr. George Lombard; (e) Source: CDC/Dr. Gilda Jones; (f) Source: CDC/Dr. Thomas F. Sellers/Emory University; (g) Source: CDC/Public Health Image Library; (h) Source: CDC/Dr. PB Smith; (i) Source: CDC - PHIL; ( j) Source: CDC; (k) Source: CDC/Dr. W.A. Clark; (l) Source: CDC/ Dr. Norman Jacobs; (m (top)) Source: CDC/Don Stalons; (m (bottom)) Source: CDC/Dr. V.R. Dowell, Jr; (n) Source: CDC

(c) Simple stain of Bacillus megaterium (1000x). Note the unstained areas within many of the cells. These are developing endospores that resist normal staining techniques and consequently appear clear after staining.

(d) Gram stain of Clostridium botulinum type A (1000x). The botulin toxin produced by this bacterium is the cause of botulism, an illness that can cause death by paralyzing the muscles of the diaphragm. The location of the developing endospores–the swelling seen at the end of some cells–is characteristic of Clostridium.

(f) Gram stain of Staphylococcus aureus (1000x). S. aureus cells are typically found in grapelike clusters. S. aureus can cause skin and bone infections, food intoxications, and a number of other diseases.

(h) Corynebacterium diphtheria, simple stain (1200x). C. diphtheria is the causative agent of diphtheria. The irregular staining seen here is characteristic of Corynebacterium.

Figure 8.5  (Continued)

(e) Gram stain of Clostridium difficile (1000x). C. difficile is a part of the gastrointestinal microbiota of most persons but can also cause Clostridium difficile disease, which is marked by watery diarrhea, nausea, and abdominal pain. Note the numerous developing endospores, which can be seen as light staining areas in many of the cells.

(g) Gram stain of Streptococcus viridins (1000x) growing in blood culture. Part of the normal microbiota, S. viridins routinely enters the bloodstream during dental procedures, presenting a risk of bacterial endocarditis to persons with certain types of cardiac aberrations. Streptococcus chains develop only in liquid cultures and increase in length as the culture ages.

(i) Gram stain of Legionella pneumophila (1000x). Gram staining reveals solitary cells and long chains of L. pneumophila, a Gramnegative bacillus responsible for legionellosis, a pneumonia beginning two to 10 days after exposure to the bacterium. L. pneumophila is also the cause of Pontiac fever, a less serious respiratory illness occurring a few hours to two days after exposure.



Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

( j) Gram stain of Bordetella pertussis (1000x). The cause of pertussis, or whooping cough, B. pertussis is a very small, encapsulated, Gram-negative coccobacillus. Pertussis is highly contagious and is most common in children and those who have not completed the primary vaccination series.

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(k) Gram stain of Acinetobacter calcoaceticus (1000x). Acinetobacter is a small, Gram-negative cell, often found in pairs, and varying in morphology from coccus to short rod. Another species in this genus, A. baumannii, is an emergent agent of disease in nosocomial settings, especially military hospitals.

(l) Gram stain of urethral exudates revealing numerous Neisseria gonorrhoeae, the cause of gonorrhea (1000x). N. gonorrhoeae is a Gram-negative coccus that, rather than being perfectly round, is flattened on one side. The cells are usually found as diplococci, with their flat sides touching.

(m) Microbiota of the oral cavity and gastrointestinal tract (Gram stain 1000x). Both Bacteroides fragilis (top) and Fusobacterium novum (bottom) are found in the oral cavity and gastrointestinal tract. Although generally regarded as commensals, both Gram-negative organisms are linked to the development of periodontal disease. (n) Gram stain of Campylobacter fetus (1000x). Campylobacter is a Gram-negative spiral rod. At least three species of Campylobacter are known to cause disease in humans.

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Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

PRE-LAB QUESTIONS Remember, Understand, Apply 1. Which of the following refers to a cellular arrangement of bacterial cells? a. rods b. coccus c. palisades d. morphology 2. A coccus that divides along two planes of symmetry would produce which arrangement? a. diplococcus b. tetrad c. coccobacillus d. streptococcus 3. Bacteria that display significant differences in shape, even within the same species, are said to be a. streptococcal. b. flexible. c. pleomorphic. d. pathogenic. 4. The primary point of staining bacterial cells is to increase the contrast between the cells and the background. a. true b. false 5. A staining technique that results in light organisms against a darkened background is a a. Gram stain. b. simple stain. c. positive stain. d. negative stain. 6. Which staining technique would provide the most accurate size of a bacterial cell? a. negative stain b. Gram stain c. simple stain d. All staining techniques would give equivalent results. 7. When preparing a bacterial smear growing on a Petri dish for staining you would a. use a loop to transfer an entire colony to the slide. b. use a needle to transfer a small amount of growth to the slide. c. use a needle to transfer a small amount of growth to a loop of water on the slide. d. use a loop to transfer a large quantity of growth to a drop of water on the slide. 8. Which of the following is a differential stain? a. simple stain b. Gram stain

c. negative stain d. All are differential stains. 9. The mordant in a Gram stain is a. crystal violet. b. Gram’s iodine. c. ethanol. d. safranin. 10. If you correctly stain a mixture of Gram-positive rods and Gram-positive cocci, you would expect to see a. purple rods and pink cocci. b. pink rods and purple cocci. c. purple rods and purple cocci. d. pink rods and pink cocci. 11. Draw and label the three most common bacterial shapes.

12. Draw and label cocci growing as diplococci, tetrads, streptococci, and staphylococci.

PERIOD ONE MATERIALS Each group should obtain: Broth cultures of: Listeria innocua Escherichia coli Slant cultures of: Bacillus megaterium Staphylococus aureus Inoculating loop and needle Methylene blue Nigrosin and/or India ink Wash bottle Bibulous paper



Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

PROCEDURE Negative Stain 1. Place a small drop of nigrosin or India ink near one end of a clean microscope slide. Use a needle to add a small amount of B. megaterium to the ink, and disperse the cells in the dye to break up any clumps. 2. Use a second slide to spread the dye across the bottom slide, as shown in Figure 8.6. 3. Allow the slide to air dry, then examine it under oil immersion. 4. Repeat this process on another slide to examine S. aureus.

Smear Preparation and Simple Stain 1. Prepare a smear of L. innocua by aseptically adding two loopfuls of the culture to the center of a clean slide. Be sure to flame sterilize your loop every time you enter the culture tube. Spread the culture over the slide, covering an area about the size of a quarter. 2. Allow the smear to air-dry completely. This should take 5–10 min. 3. Heat-fix the smear by passing the slide, with the smear on top, through the flame of your Bunsen burner three times (Figure 8.7). 4. Flood the smear with methylene blue for 1 min (Figure 8.8). 5. Gently wash the smear with water (3–4 sec).

From Liquid Media

Place a single drop of nigrosin or India ink near one end of a slide. Use a needle to disperse a small amount of organism in the dye.

From Solid Media

Two loopfuls of liquid containing organisms are placed in the center of the slide.

Culture is dispersed over middle 1/3 of slide. Place a second slide in front of the drop and move the slide backward until it touches the dye, spreading the dye across the trailing edge of the spreader slide.

Two loopfuls of water are placed in center of the slide.

A very small amount of a solid culture is dispersed with inoculating needle in water over middle 1/3 of slide.

The smear is allowed to dry at room temperature.

Slide the spreader slide forward, dragging the suspension of organisms across the slide. This should result in a smear that is quite thick at one end and very thin at the other.

After allowing the slide to air dry, it may be examined under oil immersion.

Figure 8.6  Negative-staining procedure.

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Slide is passed through flame several times to heat-kill and fix organisms to slide.

Figure 8.7  Preparation and heat fixation of bacterial smears.

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Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

Staphylococcus aureus

(1) A bacterial smear is stained with methylene blue for 1 min.

(2) Stain is briefly washed off slide with water.

Bacillus megaterium

PERIOD TWO In this period, you will perform Gram stains of each culture that was previously stained using a negative or simple stain.

MATERIALS (3) Water is carefully blotted off slide with bibulous paper.

Figure 8.8  Simple-staining procedure. 6. Blot the slide with bibulous paper to remove excess water. 7. Examine the slide under oil immersion. 8. Repeat this process on a second slide to examine E. coli.

RESULTS Make accurate drawings of each specimen, concentrating on its cellular morphology and arrangement. Escherichia coli

Each group should obtain: Broth cultures of: Listeria innocua Escherichia coli Slant cultures of: Bacillus megaterium Staphylococus aureus Two unknown cultures (these will be pure cultures of one of the four bacterial species listed here) Inoculating loop and needle Gram-staining kit (crystal violet, Gram’s iodine, 95% ethyl alcohol, safranin) Wash bottle Bibulous paper Marking pen

PROCEDURE Gram Stain 1. Label four slides with the initials of each of the bacterial species used in this exercise.

Listeria innocua The alcohol used during a Gram stain will remove any marks you made using a permanent marker. Be sure to label your slides near the edge, and watch for accidental removal of your marks.



Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

Crystal violet

30 sec

Wash

Decolorize with alcohol

8–12 sec or until solvent flows colorlessly

Wash

3 sec

Blot dry

Wash

3 sec

3 sec

Gram’s iodine

1 min

Safranin

1 min

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Figure 8.9  Gram-staining procedure.

2. Prepare a heat-fixed smear of L. innocua by aseptically adding two loopfuls of the culture to the center of a clean slide. Be sure to flame sterilize your loop every time you enter the culture tube. Spread the culture over the slide, covering an area about the size of a quarter. Prepare a heatfixed smear of E. coli in the same manner. 3. Allow the smear to air-dry completely. This should take 5–10 min. 4. Heat-fix the smear by passing the slide, with the smear on top, through the flame of your Bunsen burner three times. 5. When preparing a heat-fixed smear from bacteria growing as a solid culture, a suspension of cells is created by adding bacteria to a small drop of water on the slide. Prepare a heatfixed slide of B. megaterium by first placing a single loopful of water on a slide. 6. Use a needle to aseptically add a minute amount of bacterial growth to the water. Spread the bacterial suspension over the center of the slide, covering an area about the size of a quarter. Allow the smear to air-dry completely. This should take 5–10 min.

7. Prepare a heat-fixed smear of S. aureus in the same manner. 8. Heat fix each slide by passing it through the flame of your Bunsen burner three times. 9. Flood the slide with crystal violet for 30 sec (Figure 8.9). 10. Briefly (3–4 sec) wash the excess stain from the slide and drain off the excess water. 11. Cover the smear with Gram’s iodine for 1 min. 12. Decolorize the smear with ethyl alcohol. Allow the alcohol to flow over the slide until the runoff is colorless. This should take 8–12 sec in most cases. 13. Immediately rinse the slide thoroughly with water. This is necessary to stop the decolorizing effect of the alcohol. 14. Flood the smear with safranin for 1 min. 15. Wash gently with water to remove excess safranin. 16. Blot the slide with bibulous paper to remove excess water. 17. Examine the slide under oil immersion. 18. Repeat the Gram-staining process, being sure to properly prepare the smear, for any unknown organisms.

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Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

RESULTS Record results for each of your slides, being sure that the drawings accurately represent the cellular morphology, arrangement, and Gram reaction for each sample. Escherichia coli Staphylococcus aureus

Listeria innocua Bacillus megaterium

Gram stain each of your unknown cultures. Describe and sketch your results.

Culture # _____

Culture # _____

Can either unknown culture be conclusively identified as one of the four suspect bacteria based on the tests completed? If so, which one?

PERIOD THREE Staining of Mixed Bacterial Smears The process of Gram staining is more complex than either simple or negative staining and requires practice to achieve consistent results. Factors important to the success of a Gram stain include the following: (1) Young cultures should be used for Gram staining. Gram-positive cultures older than 24 h may sometimes stain Gram negative due to changes in the peptidoglycan layer of the cell wall that accompany aging. (2) Smears must not be too thick. Thick smears can entrap crystal violet so that it is not removed by the alcohol, leading to a false Gram-positive result. (3) Decolorization must be done for an appropriate period of time. Alcohol, if left on too long, will remove the crystal violet from Gram-positive cells, leaving them pink. Conversely, if cells are not decolorized long enough, Gram-negative cells will be seen as purple, leading to the false impression that the cells are Gram-positive. Obtaining consistent results relies primarily on decolorizing the smear for the correct period of time, which

varies somewhat from person to person. One way to determine if you are decolorizing a smear for the proper amount of time is to begin with a smear that contains two known organisms, differing in both shape and Gram reaction (Figure 8.10). The final part of this exercise will help you to fine-tune the decolorization step of your Gram stain.

MATERIALS Each student should obtain: Fresh slant culture of Bacillus megaterium Fresh broth cultures of: Staphylococcus aureus Escherichia coli Moraxella (Branhamella) catarrhalis Gram-staining kit (crystal violet, Gram’s iodine, ethyl alcohol, safranin)



Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

(a)

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(b)

Figure 8.10  Gram staining of organisms that differ in both morphology and Gram reaction can be used to perfect the Gram-staining process, preventing under- or overdecolorization of a smear. (a) Staphylococcus aureus is a Gram-positive coccus while Escherichia coli is a Gram-negative rod. (b) Bacillus megaterium is a large Gram-positive rod while Escherichia coli is a smaller Gram-negative rod. ©McGraw-Hill Education/Auburn University Research Instrumentation Facility/Michael Miller, photographer

Wash bottle Bibulous paper Glass slides Inoculating loop and needle Marking pen

(a)

EC

EC + SA

SA

BM

BM + MC

MC

PROCEDURE 1. On the left one-third of the slide, prepare a smear of E. coli. On the right one-third of the slide, prepare a smear of S. aureus. In the middle one-third of the slide, prepare a smear containing both bacteria mixed together, being careful to flame adequately to prevent contamination of the stock cultures. 2. For the second slide, prepare a smear of M. catarhalis on the left one-third of the slide, and a smear of B. megaterium on the right one-third. In the center, add a small amount of B. megaterium to two loopfuls of M catarrhalis. 3. Stain one slide at a time using the Gram stain procedure and examine with your microscope (begin with low power and work up to oil immersion). You should see organisms that differ in both shape and Gram reaction, as seen in Figure 8.11 and Table 8.1. If you do not see these results, inform your instructor, who will be able to determine what went wrong by looking at all three smears on your slides. 4. If needed, repeat the staining process with a new slide until you routinely obtain correct results. 5. Record your results in the results section.

(b)

Figure 8.11  Expected results for mixed organism Gram stains. (a) Escherichia coli cells should appear as pink rods while Staphylococcus aureus cells appear as purple cocci. If purple rods are present, underdecolorization has occurred due to not enough alcohol being used in the decolorization process while pink cocci indicate that too much alcohol was used, resulting in overdecolorization of the specimen. (b) Bacillus megaterium is Gram-positive and should appear as large purple rods while Moraxella (Branhamella) catarrhalis is a Gramnegative coccus and should appear pink. The presence of pink rods indicates overdecolorization (less alcohol should be used in the future) while the presence of purple cocci is indicative of underdecolorization and more alcohol should be used on future attempts.

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Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

TABLE 8.1

  Mixed organism Gram stain results

Slide

Correct result

Overdecolorization (too much alcohol)

Underdecolorization (too little alcohol)

E. coli / S. aureus

Pink rods, purple cocci

Pink rods, pink cocci

Purple rods, purple cocci

M. catarrhalis / B. megaterium

Pink cocci, purple rods

Pink cocci, pink rods

Purple cocci, purple rods

RESULTS Draw images from each slide, being sure to use proper colors and bacterial morphologies. E. coli plus S. aureus

B. megaterium plus M. catarrhalis

REVIEW QUESTIONS Analyze, Evaluate, Create 1. Complete the following table by providing the proper shape for each of the bacterial species listed. A look at the flowcharts in exercise 39 would be helpful. Species

Cellular morphology

Escherichia coli

Staphylococcus aureus

Listeria monocytogenes

Clostridium perfringens

2. What two staining techniques are appropriate for determining the shape and arrangement of a bacterial species, but little else?



Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

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3. Which stains can be used for negative staining, and why can these same stains not be used for simple staining?

4. What is the difference between a simple stain and a differential stain?

5. Based strictly on cellular morphology, which bacterial species used in this exercise is most easily differentiated from the others? Why?

6. What does the “Gram reaction” of a cell refer to? What color represents Gram-positive? Gram-negative?

7. Identify each of the reagents used in a Gram stain and the amount of time that each is usually left in contact with the bacterial smear. Primary stain

Mordant

Decolorizer

Counterstain

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Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

8. What is the most critical step in a Gram stain, and how can inaccuracy in this step affect your results?

9. What would happen if you inadvertently attempted to decolorize a Gram stain with water instead of ethyl alcohol?

10. Predict the color of each of the organisms at the indicated step in a properly done Gram stain. S. aureus after the mordant

E. coli after the primary stain

E. coli after the counterstain

B. megaterium after decolorization

Moraxella (Branhamella) catarrhalis after decolorization

S. aureus after the counterstain



Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

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CASE STUDY Knowing whether a cell is Gram negative or positive, and a rod or coccus is the very first step of the identification process. Often, a properly executed stain is all that is needed to rule a particular infectious agent in or out as a cause of concern. Study the details of the following case, and use your knowledge of bacterial morphology and staining procedures to answer the case study analysis questions. Identification of Bacteria Responsible for the Outbreak of Gastrointestinal Disease Late in 2011, public health officials in multiple states, with the assistance of the Centers for Disease Control and Prevention and the U.S. Food and Drug Administration, began investigating what appeared to be a food-related disease outbreak encompassing several states. Initial reports of the symptoms associated with the illness and the foods eaten by those who fell ill pointed toward one of four common bacterial species as the responsible agent. Fecal samples of ill persons as well as samples of the epidemiologically implicated foods were collected for analysis. Initial work focused on microscopic examination of the four suspect species: Clostridium perfringens, Staphylococus aureus, Escherichia coli., and Listeria monocytogenes. Public health officials worked diligently to identify the source of infections and attempted to define the scope and breadth of the epidemic by eliminating those cases of food poisoning that were clearly tied to an infectious agent different from that seen in the majority of cases. Analysis of food and fecal samples taken from ill individuals implicated Listeria monocytogenes as the infectious agent in the great majority of cases. The case was somewhat

unusual in that illnesses from Listeria are usually associated with milk, cheese, and processed meats. This outbreak was traced to cantaloupes from Jensen Farms, and investigation of the packing facilities at the farm revealed a number of instances where the bacteria could have entered and spread throughout the packing operation. One element crucial to the spread of the bacterium was the absence of an antimicrobial agent in the water that was used to wash the freshly picked fruit. Without it, the water bath used to rinse the melons became a bacterial bath, contaminating every melon that was washed in the same water. Nationwide, 147 persons reported illness, 143 of these were hospitalized, and 33 deaths were attributed to the outbreak, making it the third deadliest food-related outbreak in United States history. The actual number of listeriosis infections was assumed to be much higher, but the disease tends to be mild in most people and therefore goes unreported. Pregnant women and the elderly are at greatly increased risk from listeriosis, and in this outbreak the median age of those infected was 78, while those who died had a median age of 81. Seven cases were diagnosed in newborns or pregnant women, and one miscarriage was reported. An onslaught of medical claims and civil suits related to the outbreak led Jensen Farms to file for bankruptcy in May 2012. In October 2013, Ryan and Eric Jensen pleaded guilty to six federal misdemeanors, including introducing adulterated food into interstate commerce. Each was sentenced to 5 years probation, 6 months home detention, $150,000 in restitution, and 100 hours of community service. A civil case resulted in a confidential settlement between the victims and more than 20 of the defendants in 2015.

CASE STUDY ANALYSIS 1. In this case, which two species could not be separated on the basis of morphology or Gram reaction?

2. Bacteria within the genus Clostridium produce a protective structure called an endospore, which can be detected using a special staining technique called an endospore stain. How could an endospore stain be used to separate the two indistinguishable species?

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Case Study Exercise 8  Simple Staining, Negative Staining, and Gram Staining

3. Using the terms coccus, rod, Gram positive, Gram negative, endospore former and non-endospore former, complete the accompanying table so that each bacterial species can be properly identified.

Bacterial agent of food poisoning

Clostridium perfringens

Listeria monocytogenes

Staphylococcus aureus

Escherichia coli

REFERENCES CDC. 2012. Fatal foodborne Clostridium perfringens illness at a State Psychiatric Hospital-Louisiana, 2010. www.cdc.gov/mmwr/preview/mmwrhtml /mm6132a1.htm. CDC 2012. Outbreak of Shiga toxin-producing Escherichia coli O111 infections associated with a correctional facility dairy-Colorado, 2010. www.cdc .gov/mmwr/preview/mmwrhtml/mm6109a1.htm. CDC 2011. Multistate outbreak of listeriosis associated with Jensen Farms cantaloupe—United States, August–September 2011. www.cdc.gov/mmwr/preview/mmwrhtml/mm6039a5.htm?s_cid5mm6039a5_w. Illinois Department of Public Health. 2010. Foodborne illness linked to bakery. www.idph.state.il.us/public/press10/12.23.10Bakery.htm. Reddy, C.A., Beveridge, T.J., Breznak, J.A., Marzluf, G.A., Schmidt, T.M., and Snyder, L.R. (eds.). 2007. Methods for General and Molecular Microbiology, 3rd ed., chap. 2. Washington D.C.: ASM Press. Willey, J., Sherwood, L., and Woolverton, C. 2017. Prescott’s Microbiology, 10th ed., chap. 2. New York: McGraw-Hill.

C A S E S T U DY E X E R C I S E

9

Capsular Staining STUDENT LEARNING OUTCOME After completing this exercise, you should be able to: 1. Perform a capsule stain and differentiate between encapsulated and nonencapsulated bacterial samples.

INTRODUCTION Bacterial cells are regularly subjected to harsh environmental conditions and so are often protected by a coating of macromolecules known as a glycocalyx. Although all cells have some sort of protective layer, when the thickness, composition, and organization of this layer reach a certain level of complexity, it is accorded the status of a cellular structure and referred to as a capsule (Figure 9.1). Bacterial capsules are generally composed of repeating polysaccharide molecules, proteins, or both. The capsule adheres tightly to the cell and generally enhances the pathogenicity of bacteria such as Streptococcus pneumoniae,

Haemophilus influenzae, and Bacillus anthracis. These bacteria are able to avoid being engulfed and destroyed by white blood cells known as phagocytes, leaving them free to multiply in the tissues. Capsules also help bacteria resist dehydration, adhere to invasive devices such as catheters, and initiate the formation of biofilms (see Figure 9.1). With few exceptions, when an encapsulated bacterium loses the ability to form capsules, it also loses its pathogenicity. Because not all species of bacteria form capsules, determining that an unknown bacterial species produces a capsule can be helpful in its identification. Capsular staining is a two-step process. In the first step, an acidic dye like nigrosin or India ink is used to obliterate the background. Because these dyes are negatively charged, they will adhere to the glass of the slide but be repelled by the negatively charged bacterial cells, leaving the cells and capsules colorless against a dark background. The smear is allowed to air-dry, but there is no heat fixation because high heat can shrink the cells, resulting in a halo around the cells that could be mistaken for a capsule. In the second step, the basic dye crystal violet is used to stain the bacterial cell. Because the capsule will not stain with either dye, it is visible as a clear halo between the cell and the background. In a properly prepared capsule stain, the background will be dark while the cells themselves are purple, surrounded by a clear halo, which represents the capsule (Figure 9.2). Nonencapsulated cells will still appear as purple cells against a dark background, but the halo will be absent. Cell Capsule

Figure 9.1  Appearance of nonencapsulated Streptococcus mutans (small colonies) and encapsulated Klebsiella pneumoniae (large colonies). The capsule provides bacterial colonies with a mucoid appearance and is responsible for the bacterium’s ability to adhere to catheters and other indwelling medical devices. ©Barry Chess

Figure 9.2  Klebsiella pneumoniae display a thick bacterial capsule that can be recognized as a clear halo surrounding each darkly stained cell (1000×). ©McGraw-Hill Education/James Redfearn, photographer

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Case Study Exercise 9  Capsular Staining

PRE-LAB QUESTIONS

7. Why are bacterial smears not heat-fixed prior to capsule staining?

Remember, Understand, Apply 1. All bacterial cells produce bacterial capsules. a. true b. false 2. A capsule aides a bacterial cell by helping the bacterium to (select all that apply) a. avoid dehydration. b. avoid phagocytosis. c. adhere to surfaces. d. metabolize certain carbohydrates. 3. In a properly done capsule stain, the capsule will appear what color? a. purple b. red c. clear d. black 4. Which dye is used in the first step of a capsule stain? a. methylene blue b. safranin c. nigrosin d. malachite green 5. What would be missing when a bacterial species that does not produce a capsule is subjected to capsule staining? a. a dark background b. a purple cell body c. a clear halo 6. Name four advantages that a capsule provides a bacterial cell.

MATERIALS Each group should obtain: Skim milk slant cultures of: Klebsiella pneumoniae Enterobacter aerogenes Alcaligenes denitrificans Crystal violet stain Nigrosin or India ink Inoculating needle Marking pen

PROCEDURE 1. Label the corner of each slide with the name of the respective bacterium. 2. Place a single drop of nigrosin near one end of a slide. 3. Using an inoculating needle, aseptically transfer a very small amount of bacterial growth from a slant to the nigrosin. Use a second slide to spread the ink suspension over the slide. 4. Allow the slide to air-dry. Do not heat-fix! Heating the slide can lead to shrinkage of the cell, giving the false appearance of a bacterial capsule. 5. Place the slide on a staining rack, and flood the slide with crystal violet. Allow the smear to remain covered for 1 min. 6. Rinse the slide gently with water. 7. Blot dry with bibulous paper. 8. Examine the slide under oil immersion.

RESULTS Record your results. For each sample, be sure to indicate the cell and the presence or absence of a capsule. K. pneumoniae

E. aerogenes

A. denitrificans



Case Study Exercise 9  Capsular Staining

113

REVIEW QUESTIONS Analyze, Evaluate, Create 1. Describe and draw the way in which an encapsulated bacterium would differ from a bacterium without a capsule.

2. Which bacteria in this lab were encapsulated? Which lacked a capsule?

3. Did encapsulated bacteria appear different macroscopically (i.e., on a slant or plate) when compared to bacteria without a capsule? Explain.

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Case Study Exercise 9  Capsular Staining

CASE STUDY The following case illustrates the importance of unusual findings. Study the details of the case and answer the questions that follow. Pneumococcal Sepsis after Autosplenectomy—2005 A 15-year-old girl was admitted to the hospital after presenting at the emergency room (ER) in a semiconscious state. Feeling ill was nothing new for this patient—she had a 9-year history of systemic lupus erythematosus (SLE), a condition the ER physicians took into account as they examined her. SLE, sometimes called lupus, is an autoimmune disease in which the body produces antibodies against many of its own organs with some organs eventually becoming damaged or losing the ability to function. The specific symptoms of SLE differ, depending on which organs are affected, but kidney failure, heart problems, lung inflammation, and blood abnormalities are common. The cause of SLE is unknown. The patient’s initial workup revealed abnormally rapid breathing, fever, and low blood pressure. Additionally, her fingers and toes were cold, and she was producing no urine. The ER staff took samples of her blood and cerebrospinal fluid (CSF) and found bacteria in both. Because of the patient’s history of SLE, magnetic resonance imaging (MRI) of the abdomen was performed to assess the condition of her organs. The MRI revealed that the lupus had led to the complete destruction of the patient’s spleen, a complication called autosplenectomy that occurs in approximately 5% of SLE cases.

Asplenic individuals have low levels of both immunoglobulin M (a type of antibody) and memory B cells (a type of immune system cell that produces antibodies). Therefore, these patients are at much greater risk of infection by encapsulated bacteria. In this case, ER physicians ordered capsule staining of the bacteria isolated from the patient’s blood and CSF. Based in part on the results of the capsule staining, the bacterium isolated from both types of fluid was identified as Streptococcus pneumoniae, a heavily encapsulated species commonly encountered in asplenic patients. It is a serious sign when bacteria are found in the cerebrospinal fluid and blood as these two body compartments are generally off-limits to bacteria and have little or no normal microbial inhabitants, unlike the digestive tract or the respiratory tract. It is more difficult for microbes to enter both of these compartments for several reasons, one being that antibodies can attach to bacteria and prevent them from crossing the boundaries into these areas. Apparently this patient was missing the antibodies that would have acted against the encapsulated bacteria. The patient was treated for septic shock and respiratory failure for 9 days. Physicians administered dopamine and epinephrine to stabilize her blood pressure, as well as antibiotics to treat the underlying bacterial infection. Artificial ventilation was necessary for the first 4 days of treatment. Prior to being discharged, the patient was injected with pneumococcal vaccine and placed on prophylactic (preventive) penicillin therapy. She fully recovered.

CASE STUDY ANALYSIS 1. Define the following terms using the laboratory exercise or glossary: Biofilm Antibodies CSF

2. What is septic shock? How does sepsis figure into this case?

3. What is meant by prophylactic penicillin therapy?

REFERENCE Hühn, R., Schmeling, H., Kunze, C., and Horneff, G. 2005. Pneumococcal sepsis after autosplenectomy in a girl with systemic lupus erythematosus. Rheumatology, 44(12): 1586–1588.

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Endospore Staining STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Understand the usefulness of endospore staining. 2. Perform an endospore stain and differentiate between endospore-forming and non-endosporeforming bacterial samples.

INTRODUCTION While differential stains like the Gram stain are often used to assign a bacterial isolate to the most general of categories (Grampositive or -negative, and rod or coccus), other stains provide a means for a much more specific identification. Some bacteria have the ability to enter a resting stage, producing a tough endospore that is highly resistant to heat, cold, chemicals, and other environmental extremes that would kill a vegetative cell. Species within the genera Bacillus and Clostridium normally exist in a vegetative state, but adverse conditions, primarily a lack of carbon and nitrogen in the environment, initiate the formation

of endospores. The process takes 6–8 h, as a vegetative cell first becomes a sporangium with a developing prospore, eventually resulting in the production of a single endospore from each vegetative cell (Figure 10.1). The lifetime of an endospore is essentially limitless because the endospore will remain dormant in the environment until conditions improve, at which point the cell reenters the vegetative cycle. Often, the return of optimal growth conditions occurs when an endospore enters the body, as when Bacillus anthracis spores are inhaled. Similarly, stepping on a nail covered with Clostridium tetani spores can lead to tetanus while Clostridium botulinum endospores in food can be a source of botulism. Because so few medically important species produce endospores, determining that an unknown bacterium is an endospore former can go a long way toward providing its identification. Endospore staining is considered a structural stain because it is used to highlight a specific structure in the cell. Staining an endospore presents challenges as the protective nature of the endospore prevents the penetration of dye. Heat is used as a mordant to drive the primary stain, malachite green, into the endospore. The cell is then decolorized with water, which removes the green dye from vegetative cells but not from endospores. Finally, safranin is used to counterstain vegetative cells as well as the sporangium (but not the developing spore) of cells that are in the process of sporulation (Figure 10.2). The appearance of cells during each of these stages is summarized in Figure 10.3.

Peptidoglycan fragments

Favorable conditions Free endospore

Germination

Vegetative Cycle

Binary fission

Sporulation Cycle

Mature endospore

Sporangium with developing prospore

Figure 10.1  Vegetative and sporulation cycles in endospore-forming bacteria. The sporulation cycle begins (counterclockwise from center) with the replication of the bacterial chromosome (black) and the appearance of the developing prospore (green). The prospore contains a copy of the bacterial chromosome along with the minimum structures and chemicals necessary for survival. The surrounding sporangium continues to manufacture compounds required for endospore formation, eventually lysing when the process is complete. The free endospore may remain dormant for thousands of years until favorable environmental conditions cause it to germinate and reenter the vegetative cycle. 115

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Case Study Exercise 10  Endospore Staining Endospore former

Reagent

Non-endospore former

None (heat-fixed smear)

Malachite green and heat

Water

Safranin

Figure 10.2  Endospore stain of Bacillus sp. Mature endospores stain green while vegetative cells stain pink. Pink cells with a green center are sporangia with developing endospores (1000x). Source: CDC/

Figure 10.3  Color changes that occur at each step of the endospore-staining process.

Larry Stauffer, Oregon State Public Health Laboratory

PRE-LAB QUESTIONS Remember, Understand, Apply 1. Endospores are produced by bacteria in the genera (choose all that apply) a. Bacillus. b. Staphylococcus. c. Klebsiella. d. Clostridium. 2. Endospores are usually formed in response to a. high temperatures. b. a lack of carbon and nitrogen. c. antibiotics. d. a lack of oxygen. 3. In a properly done endospore stain, the endospore will appear what color? a. purple b. pink c. green d. clear

Bacterial species Staphylococcus aureus Escherichia coli Bacillus anthracis

Gram reaction  (+/−)

4. Which dye is used in the first step of an endospore stain? a. methylene blue b. safranin c. nigrosin d. malachite green 5. An endospore stain is an example of a a. simple stain. b. negative stain. c. structural stain. d. differential stain. 6. Name three bacterial pathogens that form endospores as well as the disease associated with each.

7. Complete the table:

Gram stain color

Endospore  (+/−)

Endospore stain color



Case Study Exercise 10  Endospore Staining

MATERIALS Each student or group should obtain: 48–72 h nutrient agar culture of Bacillus megaterium, B. subtilis, or B. cereus Electric hot plate and small (25 ml) beaker (Schaeffer-Fulton method only) Microwave oven and empty Petri dish (microwave method only) Endospore-staining kit (5% malachite green, safranin) Water bottle Glass slides Bibulous paper Inoculating loop and needle Marking pen

PROCEDURE Schaeffer-Fulton Method This procedure is illustrated in Figure 10.4. 1. Label a slide in the upper left-hand corner for identification, and prepare a heat-fixed smear of the appropriate culture. 2. Saturate the smear with malachite green, and steam the slide over boiling water for 5 min. Do not allow the stain to dry on the smear. Add additional stain if required to keep the smear wet during the heating process. 3. Allow the slide to cool slightly so that it will not crack. Rinse with water for 30 sec. 4. Counterstain with safranin for 30 sec. 5. Rinse briefly to remove excess safranin.

(1) Cover the heat-fixed smear with malachite green. Steam over boiling water for 5 min. Add additional stain if smear begins to dry.

(3) Counterstain with safranin for about 30 sec.

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6. Blot dry with bibulous paper. 7. Examine the stained specimen under low power, high power, and oil immersion. Record your observations in the Results section of this exercise. Microwave Method 1. Label a slide in the upper left-hand corner for identification, and prepare a heat-fixed smear of the appropriate culture. 2. Place a cut-to-fit piece of paper towel (two layers of paper) into the bottom of an empty Petri dish. Saturate the toweling with water. 3. Place the heat-fixed slide on top of the paper towel, and flood the slide with malachite green. 4. Place the slide in the microwave, and heat for 30 sec at full power. 5. Remove the Petri dish from the microwave, and allow the slide to cool slightly so that it will not crack (30 sec is adequate). Rinse with water for 30 sec. 6. Counterstain with safranin for 30 sec. 7. Rinse briefly to remove excess safranin. 8. Blot dry with bibulous paper. 9. Examine the stained specimen under low power, high power, and oil immersion. Record your observations in the Results section of this exercise. Slides that have been stained for endospores will generally have large patches of dried malachite green stain on them. Don’t mistake these blobs for the (much smaller) endospores.

(2) After the slide has cooled sufficiently, rinse with water for 30 sec.

(4) Rinse briefly with water to remove safranin.

Figure 10.4  Procedure for the Schaeffer-Fulton method of endospore staining.

(5) Blot dry with bibulous paper and examine slide under oil immersion.

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Case Study Exercise 10  Endospore Staining

RESULTS Draw images from your slide, being sure to use proper colors and bacterial morphologies.

REVIEW QUESTIONS Analyze, Evaluate, Create 1. What is the advantage of performing a Gram stain prior to an endospore stain?

2. Why are older cultures typically used for endospore staining? (A hint: Why would a fresh young culture be inappropriate for endospore staining?)

3. After the specimen is stained with malachite green, the slide is washed for 30 sec. In most other staining techniques, washes are only a few seconds in length. Why is such a long rinse used in this case?

4. What would you observe if you performed an endospore stain on a culture of Bacillus megaterium but neglected to apply any heat during the staining process?



Case Study Exercise 10  Endospore Staining

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CASE STUDY Mention anthrax and everyone’s first thought goes to terrorism. For those with a little more knowledge of microbiology, second thoughts tend to center on ranchers, sheepherders, and leather workers. But the most recent cases of anthrax have come from a decidedly different source. Read the following case and use your knowledge of endospore-forming organisms to answer the case study analysis questions. Drumroll please . . . Inhalation Anthrax Associated with Dried Animal Hides—London, 2008 A 35-year-old male presented to a London hospital complaining of difficulty breathing. His symptoms progressed quickly, and he was transferred to the hospital’s intensive treatment unit suffering from respiratory failure, which soon progressed to multiple organ failure. A blood culture revealed Gram-positive, encapsulated, nonmotile rods preliminarily identified as Bacillus anthracis. This is the bacterium that causes the disease anthrax, and it has the ability to survive for long periods of time without water or nutrition. The presence of B. anthracis was later confirmed by the Novel and Dangerous Pathogens Division of Britain’s Health Protection Agency. The patient was a drum maker by trade, handcrafting traditional African drums from dried animal hides. This process required him to soak the hides in water for an hour and then scrape the hair off with a razor, thereby releasing large quantities of dust into his studio. Most of the animal skins were

goat hides imported from Gambia. Investigators from the Health Protection Agency examined the drum maker’s property for the presence of the anthrax bacterium. In his studio, they found endospores of B. anthracis on one of five drums and on a few animal skins. No other traces of the bacterium were found at the property. Although rare in the U.K.—and in the United States, for that matter—B. anthracis is found throughout much of the world. Its ability to form endospores and survive harsh environmental conditions (years of heat, cold, and ultraviolet radiation, along with a complete lack of water or nutrients) ensures that many endospores are found in soil. As animals graze, lie, or roll on the ground, some of the endospores are transferred onto their bodies. Livestock in Africa, Asia, and the Middle East account for the majority of anthrax cases seen worldwide. Despite treatment with rifampin, ciprofloxacin, and clindamycin, as well as with anti-anthrax immunoglobulin, the drum maker died about 2 weeks later. Postexposure prophylaxis was given to eight persons, including the patient’s immediate family, the main supplier of the skins, a person who assisted with the drum making, and a hospital worker. This incident was very similar to two 2006 cases in which drum makers in New York City and Scotland contracted anthrax while scraping animal hides for drumheads. In all three cases, the hides were imported from Africa, where anthrax is endemic.

CASE STUDY ANALYSIS 1. Cowboys, ranch hands, shepherds, and persons who work tanning leather are all at increased risk for being infected with anthrax. Why is this?

2. When untreated, anthrax is virtually always a fatal disease. The patient in this case had many social contacts yet only eight people were prescribed prophylactic antibiotics. Why were more people not treated with antibiotics?

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Case Study Exercise 10  Endospore Staining

THERE’S MORE TO THE STORY . . . As seen in this case, endospores may be found on animal hides, and anthrax itself is even known as Woolsorter’s disease because sheep often harbor Bacillus anthracis endospores in their coat. Whether wearing a respirator to protect against the inhalation of animal dander or receiving a tetanus shot after stepping on a nail, it doesn’t take a medical professional to recognize that endospores are widespread throughout the environment. In fact, isolation and characterization of Bacillus species is a fairly straightforward process that your instructor may have you carry out. A note of caution: You will be working with wild microorganisms, whose virulence and pathogenicity are completely unknown. Be sure to employ proper microbiological safety techniques. Aside from the first two steps, what follows is merely the outline of a full protocol for the isolation and identification of Bacillus species. You will need to develop a step-by-step protocol, using your lab book and any other resources you have available before you begin. 1. Add a pinch of dirt, dust, or decaying material to a small glass culture tube containing one ml of distilled water. 2. Heat the tube in a water bath at 80°C for 15 min. This will enrich the sample for endospore formers.

3. Plate your sample onto nutrient agar and incubate for 24 h. Besides forming endospores, most species in the genus Bacillus are Grampositive rods that produce catalase and grow as aerobes or facultative anaerobes. Test several colonies—remember to do simple tests first—to determine which of your isolates may fit this description. Pick one or two colonies to culture to a new plate and continue with your identification. Some, though by no means all, exercises in your lab book that may prove useful include ∙∙ Exercise 8, Simple Staining, Negative Staining, and Gram Staining ∙∙ Exercise 10, Endospore Staining ∙∙ Exercise 39, Identification of Bacterial Unknowns ∙∙ Exercise 45, Streak Plate Isolation ∙∙ Exercise 48, Fluid Thioglycollate Medium ∙∙ Exercise 68, Catalase Test Be sure to adequately describe your isolate, including its macroscopic (colonial) and microscopic (cellular) appearance, along with its physiological and biochemical characteristics. When finished, ensure that you have properly disposed of all biohazardous waste materials.

REFERENCE CDC. Anthrax. http://www.cdc.gov/anthrax.

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11

Acid-Fast Staining STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Understand the usefulness of acid-fast staining. 2. Perform an acid-fast stain and differentiate between acid fast and non-acid-fast bacterial samples.

INTRODUCTION The acid-fast stain is a differential stain primarily used to detect members of the genus Mycobacterium, including the pathogens M. tuberculosis and M. leprae, the causative agents of tuberculosis (TB) and leprosy, respectively. In addition, bacteria in the genus Nocardia and even some protozoan parasites (Cryptosporidium and Isospora) show at least some degree of acid-fast behavior. Because so few organisms are acid-fast, the stain is generally only used when infection by an acid-fast organism is suspected. Acid-fast staining of sputum samples often directs treatment, allowing the physician to administer drugs such as isoniazid and rifampin, which are commonly prescribed for TB infection, long before these slow-growing bacteria could be cultured in the laboratory. Acid-fast bacteria contain within their cell walls a waxy material called mycolic acid, which prevents most stains from penetrating the cell, leading to a light purple color when Gram stained. However, when heat is used to soften the mycolic acid, the primary stain, carbol-fuschin, enters the cells and is not removed when decolorized with acid-alcohol (3% hydrochloric acid in 95% ethanol), leaving the cells a deep red color (Figure 11.1). Cells that are non-acid-fast are easily decolorized by acid-alcohol, leaving them colorless until the counterstain, methylene blue, is applied. The differences in the appearance of acid-fast and non-acid-fast cells at each step of the staining process are seen in Figure 11.2.

Figure 11.1  Acid-fast stain of Mycobacterium smegmatis (red, acidfast rods) and Staphylococcus aureus (blue, non-acid-fast cocci). ©The McGraw-Hill Companies, Inc./Auburn University Research Instrumentation Facility/Michael Miller, photographer

Reagent

Acid-fast

Non-acid-fast

None (heat-fixed smear) Carbolfuschin and heat

Acid-alcohol

Methylene blue

Figure 11.2  Color changes that occur at each step of the acid-fast staining process.

PRE-LAB QUESTIONS Remember, Understand, Apply 1. Which genus contains acid-fast bacteria? a. Bacillus b. Staphylococcus c. Streptococcus d. Mycobacterium

2. What component of the cell wall is responsible for the manner in which acid-fast bacteria absorb and release stain? a. isoniazid b. mycolic acid c. lipopolysaccharide d. peptidoglycan

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Case Study Exercise 11  Acid-Fast Staining

3. In a properly done acid-fast stain, non-acid-fast bacteria will appear what color? a. purple b. pink c. green d. blue 4. Which disease is caused by members of the genus Mycobacterium? a. cholera b. diabetes c. leprosy d. tetanus

5. An acid-fast stain is an example of a a. simple stain. b. negative stain. c. structural stain. d. differential stain. 6. Define the following terms using the laboratory exercise or glossary: Differential stain

7. Complete the table: Bacterial species

Gram reaction (+/−)

Gram stain color

Acid-fast (+/−)

Acid-fast stain color

Staphylococcus aureus Mycobacterium tuberculosis Escherichia coli

MATERIALS Each student or group should obtain: A nutrient agar slant culture of Mycobacterium smegmatis A nutrient broth culture of Staphylococcus aureus Acid-fast staining kit (carbol-fuschin, acid-alcohol, methylene blue) Inoculating loop and needle Marking pen For Ziehl-Neelson staining procedure: Electric hot plate 50 ml beaker For microwave procedure: Microwave oven Empty plastic Petri dish

PROCEDURE Two different acid-fast staining procedures are provided. Your instructor will inform you as to which one to use. Ziehl-Neelsen Method This procedure is illustrated in Figure 11.3. 1. Label a slide in the upper left-hand corner for identification. 2. Prepare a mixed smear by transferring two loopfuls of S. aureus to a slide and adding to it a very small amount of M. smegmatis. The acid-fast bacteria tend to form clumps that will have to be broken apart with the inoculating needle. Air dry and heat-fix the slide.

3. Saturate the smear with carbol-fuschin, and steam the slide over boiling water for 5 min. Add additional stain if required to keep the smear from drying out. 4. Allow the slide to cool slightly so that it will not crack. Rinse with acid-alcohol for 8–12 sec. 5. Rinse briefly with water to stop the decolorizing effect of the acid-alcohol. 6. Counterstain with methylene blue for 30 sec. 7. Rinse briefly to remove excess methylene blue. 8. Blot dry with bibulous paper. 9. Examine the stained specimen under low power, high power, and oil immersion. Record your observations in the Results section of this exercise.

Even when examining an acid-fast culture, only a few cells are likely to appear red. In a clinical lab, to classify a sample as acid-fast negative, 300 visual fields must be examined without detecting any acid-fast cells.

Microwave Method 1. Label a slide in the upper left-hand corner for identification. 2. Prepare a mixed smear by transferring two loopfuls of S. aureus to a slide and adding to it a very small amount of M. smegmatis. The acid-fast bacteria tend to form clumps that will have to be broken apart with the inoculating needle. Air dry and heat-fix the slide.



Case Study Exercise 11  Acid-Fast Staining

(1) Cover smear with carbolfuchsin. Steam over boiling water 5 min. Add additional stain if stain boils off.

(2) After slide has cooled, decolorize with acid-alcohol for 8–12 sec.

(3) Stop decolorization action of acidalcohol by rinsing briefly with water.

(4) Counterstain with methylene blue for 30 sec.

(5) Rinse briefly with water to remove excess methylene blue.

(6) Blot dry with bibulous paper. Examine directly under oil immersion.

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Figure 11.3  Ziehl-Neelsen acid-fast-staining procedure. 3. Place a cut-to-fit piece of paper towel (two layers of paper) into the bottom of an empty Petri dish. Saturate the toweling with water. 4. Place the heat-fixed slide on top of the paper towel, and flood the slide with carbol-fuschin. 5. Place the cover on the dish and heat in the microwave for 30 sec at full power. 6. Remove the Petri dish from the microwave, and allow the slide to cool slightly so that it will not crack. Decolorize with acid-alcohol for 8–12 sec.

7. Rinse briefly with water to stop the decolorizing effect of acid-alcohol. 8. Counterstain with methylene blue for 30 sec. 9. Rinse briefly to remove excess methylene blue. 10. Blot dry with bibulous paper. 11. Examine the stained specimen under low power, high power, and oil immersion. Record your observations in the Results section of this exercise.

RESULTS Draw images from your slide, being sure to use proper colors and bacterial morphologies.

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Case Study Exercise 11  Acid-Fast Staining

REVIEW QUESTIONS Analyze, Evaluate, Create 1. List each of the reagents used in an acid-fast stain, along with its purpose in the staining process.

2. Why is an acid-fast stain used so much less frequently than a Gram stain?

CASE STUDY Both tuberculosis and leprosy are, fortunately, exceedingly rare in the United States. However, a number of other species in the genus Mycobacterium are associated with disease. Study the following case and use your knowledge of acid-fast bacteria to answer the case study analysis questions. Tattoo-Associated Nontuberculous Mycobacterial Skin Infections — New York, 2011–2012 On January 4, 2012, the Monroe County (New York) Department of Public Health received a report of a person with a persistent rash associated with a new tattoo. The rash first appeared in October 2011, one week after receiving the tattoo, and on closer inspection was found to be a bacterial infection of the skin and underlying soft tissue, with the infection limited to those areas of the skin that were tattooed with gray ink. A skin biopsy was performed, and bacteria isolated from the infection stained weakly Gram positive. Additional staining showed the isolate to have an acid-fast cell wall, presumptively identifying it as a member of the genus Mycobacterium. Biochemical and physiological testing confirmed the identity of the infectious agent as Mycobacterium chelonae, a species known to cause nontuberculous mycobacterial (NTM) skin and soft tissue infections that can range from mild inflammation to severe abscesses requiring extensive surgical debridement. The infections are difficult to combat, requiring a minimum of four months of treatment with a combination of two or more antibiotics. The tattoo artist stated that he had been using the same brand of ink

since May of the previous year. Using a list of his customers, a total of 19 infections were identified, including 14 in which M. chelonae was isolated from the infection. Mycobacterium chelonae and a related species, Mycobacterium abscessus, are found in water, so the addition of nonsterile water to ink during its manufacture or at its point of use could lead to contamination and potentially result in infection. The artist in this case specified that he bought his ink prediluted from the manufacturer and added no water himself. Investigators isolated M. chelonae from tissue specimens and from one opened and one unopened bottle of gray ink. A review of the tattoo artist’s practices revealed no other potential sources of contamination, and examination of the ink manufacturer’s facility revealed no ongoing contamination. Under the Federal Food, Drug, and Cosmetic Act, tattoo inks are considered to be cosmetics, and there are no specific FDA regulations requiring tattoo inks to be sterile. However, the CDC recommends that ink manufacturers ensure ink is sterile and that tattoo artists avoid contamination of ink through dilution with nonsterile water. Beyond the risk of initial contamination, dilution of ink with water will dilute any preservatives in the ink, reducing their effectiveness. While no federal laws regulate the practice of tattooing, in some areas local authorities require bloodborne pathogens training and the use of hygienic practice during tattooing. Certain jurisdictions, such as Los Angeles County, go one step further and require that sterile water be used in tattoo ink dilution.



Case Study Exercise 11  Acid-Fast Staining

CASE STUDY ANALYSIS 1. In the case seen here, little thought was given to examining family members of patients for Mycobacterium chelonae or Mycobacterium abscessus infection. By contrast, infection with Mycobacterium tuberculosis usually leads to an immediate public health investigation. Why are these organisms treated so differently despite being in the same genus? (You may have to research M. tuberculosis to frame your answer.)

2. Treatment of mycobacterial infections generally entails a four- to nine-month antibiotic regimen. How is the length of treatment connected to the fact that mycobacterial species tend to grow very slowly?

REFERENCES CDC. 2012. Tattoo-Associated Nontuberculous Mycobacterial Skin Infections-Multiple States, 2011–2012. Morbidity and Mortality Weekly Report, 61: 653–656. CDC. Division of Tuberculosis Elimination (DTBE). http://www.cdc.gov/tb/. Tille, P.M. 2017. Bailey and Scott’s Diagnostic Microbiology, 14th ed., p. 538. St. Louis: Elsevier. Reddy, C.A., Beveridge, T.J., Breznak, J.A., Marzluf, G.A., Schmidt, T.M., and Snyder, L.R. (eds.). 2007. Methods for General and Molecular Microbiology, 3rd ed., chap. 2. Washington D.C.: ASM Press.

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NOTES

C A S E S T U DY E X E R C I S E

12

Viable Plate Count STUDENT LEARNING OUTCOMES After completing this exercise, you should be able to: 1. Determine the number of bacteria in a liquid sample using a viable plate count. 2. Calculate and carry out dilutions of a liquid sample.

INTRODUCTION The testing of liquids for the presence of microbial contaminants is just part of a routine day for thousands of microbiologists. Whether analyzing drinking water, orange juice, or buffers used to dilute pharmaceutical products, the goal is the same, ensuring that living microbes in a sample are below a set limit. The testing of milk serves as a common example of this process. As a bodily fluid, milk in the udder of a cow is sterile. During the collection process, however, it is inoculated with the normal microbiota of the cow. Because milk has such a rich composition, being laden with water, proteins, fats, and hormones, microbial growth is a common occurrence. Since milk is such an integral part of so many people’s lives and since its consumption can entail very real risks, the entire production process, from cow to cup, is tightly regulated. The main safeguard against bacterial infections acquired from milk is pasteurization. In this process, heat is used to kill many bacterial species, resulting in a product that, while not sterile, has a reduced microbial load. This decrease in bacterial diversity and number results in a product that is safer to drink and has a longer shelf life. Raw milk refers to milk that has not been pasteurized as part of its processing. Dairies that sell raw milk rely on more stringent standards of cow health as well as rigorous collection standards. Despite these efforts, raw milk remains a relatively common source of Salmonella and Escherichia coli outbreaks. For both raw and pasteurized milk, the Food and Drug Administration has set limits on the number of both coliforms and total bacteria that may be present in milk before and after processing and/or pasteurization. Technically speaking, coliform bacteria are Gramnegative rods that ferment the sugar lactose, producing acid and gas. The name itself refers to the fact that these bacteria look and act like a “form” of E. coli. Less technically speaking, coliforms— especially E. coli—are common inhabitants of the gastrointestinal tract, and their presence in food or water is an indicator of fecal contamination. Because food, milk, and water are so commonly

Figure 12.1  When grown on MacConkey agar, lactose-fermenting bacteria should be deep pink or red in color while lactose nonfermenters are white to colorless. Because MacConkey agar selects only Gram-negative bacteria for growth, red colonies can presumptively be considered coliforms (Gram-negative lactose fermenters) while white or clear colonies are considered noncoliforms. ©Barry Chess

tested for the presence of coliforms, several types of selective and/or differential media have been formulated to make testing as simple as possible. One commonly used medium, Mac­Conkey agar, contains lactose, along with a pH indicator that turns bacterial colonies pink if the pH of the medium drops, as would happen if a coliform bacterium were fermenting lactose. Nonfermenters of lactose do not cause the pH to drop and consequently produce colonies that are clear or white (Figure 12.1). Bacterial counts in milk are often determined using a viable plate count, both as part of routine testing and whenever milk products are thought to be the source of a disease cluster. In this method, a sample of milk to be counted is serially diluted to produce several samples with decreasing cell densities. Aliquots of the dilutions are then plated onto media, and the colonies produced are counted after incubation. Once the number of colonies is known, it is a simple matter to mathematically calculate the number of cells in the original sample. Each dilution is represented by a dilution factor, a term that corresponds to the amount of the original sample still present in the current sample. For example, if 1 ml of a sample is diluted into 9 ml of diluent (the fluid used for dilution), so that the original sample makes up 1/10 of the current volume, then the dilution factor is 10. If 1 ml were diluted in 99 ml, the dilution factor would be 100 because the sample would be 1/100 as concentrated,

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Case Study Exercise 12  Viable Plate Count

and so on. The use of dilution factors, along with careful measurement, allows the accurate determination of cell densities in any liquid. When several dilutions are performed in sequence, a serial dilution is the end result, with each sample being less concentrated than the one before. The most important part of performing a serial dilution is being able to calculate exactly how dilute a sample is when compared to the original. This can easily be done by remembering two facts. 1. A dilution factor is always found by dividing the volume of the first sample by the volume of the first sample plus the volume of the diluent, and then taking the reciprocal. For example, diluting 1 ml of a milk sample into 99 ml of sterile water results in a 1/100 dilution and a dilution factor of 100. 2. In a serial dilution, the final dilution factor is simply the product of each individual dilution factor. For example, if an undiluted liquid (dilution factor of 1) is diluted into 9 ml of diluent, 1 ml of this sample is diluted into 9 ml of diluent, and 1 ml of this sample is diluted into 9 ml of diluent, then the final dilution factor is (1)(10)(10) (10) = 1000. Finally, the cell density of the original broth is a function of the volume of the sample plated onto the media, the dilution of the sample, and the number of colonies on the plate. The original cell density (OCD) can be calculated using the formula: OCD =

(colonies on plate) (volume of sample plated)

× dilution factor

2. Raw milk is commonly associated with outbreaks of which bacteria? a. Salmonella and Escherichia coli b. Staphylococcus aureus and Streptococcus pyogenes c. Pseudomonas aeruginosa and Enterococcus faecalis d. Bacillus anthracis and Mycobacterium tuberculosis 3. MacConkey agar selects for the growth of which type of organism? a. Gram-negative b. Gram-positive 4. Two ml of milk are diluted into 8 ml of sterile water. What is the dilution factor? a. 10 b. 8 c. 5 d. ¼ 5. A plate inoculated with 0.1 ml of a 1/100 dilution of milk produced 30 colonies. How many organisms are in 1 ml of the undiluted sample? a. 30 b. 300 c. 3000 d. 30,000 6. Define the following terms using the laboratory exercise or the glossary: Viable

For example, if 0.1 ml from a sample with a dilution factor of 10,000 (104) is used to inoculate a plate and 122 colonies are counted after incubation, the OCD is: OCD =

122 colonies × 10,000 0.1 ml

Coliform

OCD = 1.2 × 107 colonies/ml One last point. Although we tend to think of a colony arising from a single cell, it is important to remember that for some organisms, single cells are rarely if ever seen. The term colony forming unit (CFU) allows us to sidestep the problem of exactly how many cells were involved in the formation of a single colony by grouping together pairs, tetrads, clusters, and chains under a common term. The fact remains that a colony arises from the smallest unit of colony formation, be it one cell, two, four, or more. For the sake of accuracy, therefore, the term CFU is preferable to cells and will be used from this point forward.

Differential media

Colony forming unit

PRE-LAB QUESTIONS Remember, Understand, Apply 1. Once milk has gone through the process of pasteurization it is considered sterile. a. true b. false

7. For each of the media in the accompanying table, indicate what type of bacteria is selected for and how lactosefermenting bacteria are differentiated from lactosenonfermenting bacteria. (Referring to the exercise in the lab manual associated with each media type will certainly help.)



Case Study Exercise 12  Viable Plate Count

Specificity of selection

Medium

Color of lactose-fermenting colonies

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Color of lactose-nonfermenting colonies

MacConkey agar Eosin methylene blue agar Hektoen enteric agar

PERIOD ONE The technique used in this exercise is a slightly simplified version of the protocol required by the FDA. Each student team will be responsible for determining the bacterial count of coliforms and total bacteria from one type of milk, either raw or pasteurized.

MATERIALS Each group should obtain: Three plates of nutrient agar* Three plates of MacConkey agar* Three tubes each containing 9 ml of sterile water (blanks) One screw top tube containing pasteurized milk (odd-numbered student teams) One screw-top tube containing raw milk (even-numbered student teams) Six serological pipettes, 1 ml Small (100 ml) beaker containing 50 ml of ethanol Glass spreading rod Large beaker that fits over the smaller beaker Marking pen * For best results, these plates should be allowed to dry at room temperature for several days, or overnight at 37°C prior to being inoculated.

PROCEDURE 1. The dilution and inoculation procedure is illustrated in Figure 12.2 and Table 12.1. 2. Label the bottom of all 6 plates and the 3 water blanks with your name and lab time. 3. Label the water blanks 1/10, 1/100, and 1/1000. 4. After making sure the sample container is completely closed, vigorously shake your milk sample for 10 sec. 5. Within 3 min, aseptically transfer 1 ml of milk to the first (1/10) tube. 6. Pipette up and down several times to mix, and transfer 1 ml of the 1/10 dilution to the 1/100 tube. 7. Pipette up and down several times to mix, and transfer 1 ml of the 1/100 dilution to the 1/1000 tube. Pipette the

sample in the 1/1000 tube up and down several times to mix. 8. Inoculate the nutrient agar plates with the 1/10, 1/100, and 1/1000 dilutions. Inoculate each plate with 0.5 ml of the appropriate dilution, and spread the inoculum with a sterile glass spreader. Do not invert the plates for at least 10 min. 9. Label the MacConkey agar plates as undiluted, 1/10, and 1/100. Inoculate the first plate with undiluted milk (shake the milk again if it has been sitting more than 3 min since last being mixed), the second plate with the 1/10 dilution, and the third plate with the 1/100 dilution. Spread the inoculum on each plate using a sterile glass spreader. 10. Incubate the plates at 32°C for 24–48 h. Be sure to properly dispose of all tubes, pipettes, and other equipment.

TABLE 12.1

  Inoculating a Plate with a Glass Spreader

Occasionally, bacteria may be spread across the surface of a plate using a sterile glass rod bent into the shape of a hockey stick. The rod is sterilized by dipping it in ethanol and then passing it through the flame of a Bunsen burner. Once the alcohol has burned off, the spreader is ready to use. The following protocol explains how to use the spreader to evenly distribute bacteria after they’ve been added to the plate. 1. Place the spreader rod in the beaker, and add enough ethanol to the beaker to cover the lower portion of the rod. 2. Arrange the items on your bench so that the Bunsen burner is between the alcohol and the Petri dish. This reduces the chance of accidentally setting the alcohol aflame. Have nearby an empty beaker large enough to place over the small beaker and spreading rod. If the alcohol catches fire, simply place the large beaker over the smaller beaker to extinguish the flame. 3. Holding the upper end of the spreader rod, remove it from the alcohol and pass it through the flame of the Bunsen burner, allowing the alcohol to ignite. Be sure to keep your hand above the spreader so that the flaming alcohol doesn’t run onto your hand. Wait until the alcohol has completely burned off before continuing. 4. Place the lower portion of the spreader flat against the agar plate. Rotate the plate using your thumb and middle finger while moving the spreader back and forth. If a turntable is used, gently spin the turntable while moving the spreader back and forth. In either case, finish by rotating the plate one complete revolution while holding the spreader against the edge of the plate. 5. Return the spreader to the alcohol. There is no need to flame it again.

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Case Study Exercise 12  Viable Plate Count 1 ml

1 ml

Milk sample

0.5 ml

1 ml

1/10 dilution

0.5 ml

1/100 dilution

0.5 ml

0.5 ml

0.5 ml

Nutrient agar

MacConkey agar

1/1000 dilution

Nutrient agar

0.5 ml

Nutrient agar

MacConkey agar

MacConkey agar

Figure 12.2  Dilution and inoculation procedure for a viable plate count. plates containing between 25 and 250 colonies. Record your results. 4. Compare the growth between plates inoculated with each type of milk (raw versus pasteurized). Using the morphological appearance of the colonies on each plate, determine the number of different types of colonies present on each plate. Review Exercise 41 for help in differentiating colony morphology.

PERIOD TWO PROCEDURE 1. Retrieve your plates from the incubator. The growth of coliforms and noncoliforms on MacConkey agar is illustrated in Figure 12.1. 2. Use a colony counter to count the number of colonies on each plate. Any plate with fewer than 25 colonies should be recorded as TFTC (too few to count) while any plate with more than 250 colonies as TMTC (too many to count). Count colonies of all sizes. 3. Use dilution factors to establish the total number of each type of microorganism per milliliter of milk, using only those

Distinct colony types Raw milk Pasteurized milk

Raw milk Type of media used Total bacterial count Gram-negative bacteria count Coliform count

Undiluted

1/10

1/100

Dilution factor

Organisms/ml



Case Study Exercise 12  Viable Plate Count

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Pasteurized Milk Type of media used

1/10

1/100

1/1000

Dilution factor

Organisms/ml

Total bacterial count Gram-negative bacteria count Coliform count

REVIEW QUESTIONS Analyze, Evaluate, Create 1. The FDA requires that Grade A pasteurized milk have no more than 20,000 CFUs per ml. Based on this standard, what is the maximum number of CFUs that are permissible in: 0.1 ml?

0.01 ml?

0.001 ml?

2. Why was the milk used for coliform counting diluted to a lesser extent than the milk used for total bacterial counting?

3. What is the relationship between a cell, a CFU, and a colony?

4. FDA standards call for counting only plates with between 25 and 250 colonies per plate. Those plates with fewer than 25 colonies are reported as TFTC (too few to count), and those with more than 250 as TMTC (too many to count). Why do you think the FDA has instituted this recommendation?

5. Which milk, raw or pasteurized, had a greater variety (not number) of colonies? Explain this fact.

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Case Study Exercise 12  Viable Plate Count

CASE STUDY The Centers for Disease Control and Prevention put it very plainly, “Raw milk can carry harmful bacteria and other germs that can make you very sick or kill you.” Yet, many people prefer to drink raw [unpasteurized] milk for supposed health benefits, most of which simply don’t exist. Study the following case, and then answer the case study analysis questions. Campylobacter jejuni Infections Associated with Raw Milk Consumption—Utah, 2014 In May 2014, the Utah Public Health Laboratory (UPHL) identified three patients suffering from infection with Campylobacter jejuni, a Gram-negative bacterium that is one of the most common causes of diarrheal illness in the United States. The three samples had identical DNA profiles, indicating a common source, and the UPHL informed the Utah Department of Health (UDOH) of a probable outbreak. In most cases, campylobacteriosis (infection with Campylobacter) causes diarrhea, abdominal cramping, fever, nausea, and vomiting lasting about a week. Rarely, especially when the person involved has a weakened immune system, Campylobacter may spread to the bloodstream and cause a lifethreatening infection. All three persons had consumed raw (unpasteurized) milk from Ropelato Dairy in northern Utah. The first two patients were a parent and child whose symptoms began on May 10. Both were hospitalized, and the parent died a week later of multisystem organ failure. The third patient’s symptoms began on May 11. Over the next 6 months, a total of 99 cases of C. jejuni infection were identified, 10 patients were hospitalized, and 1 died. Among the 98 persons interviewed, 52 reported drinking raw milk from Ropelata Dairy. Among 16

the 41 persons who reported not drinking raw milk, just over half reported eating queso fresco, a soft cheese that, when made from raw milk, supports the growth of bacteria. Inspectors from the Utah Department of Agriculture and Food (UDAF) regularly conduct dairy inspections, and Ropleto Dairy had been inspected on June 1, prior to the UDAF being apprised of the outbreak. In addition to ensuring that the dairy follows proper safety procedures, samples of milk are tested to determine the total number of bacteria, number of coliform bacteria, and number of somatic cells they contain. Coliforms are a group of bacteria resembling E. coli in their physiological behavior, and their presence is indicative of fecal contamination. Although Campylobacter is commonly found in feces, it is not a coliform, and tests designed to detect coliforms will not identify its presence. Somatic cells are cells present in the milk that are from the cow itself. Mostly leukocytes (white blood cells) that are produced in response to infection, a high number of somatic cells is indicative of a sick cow. To be sold in Utah, raw milk may contain no more than 20,000 bacterial cells, 10 coliforms, and 400,000 somatic cells per milliliter. Ropelata Dairy met this standard on their June 1 inspection, as well as two inspections on June 12 and July 13 ordered after the outbreak was recognized. Based on their successful test results, the dairy was allowed to continue selling raw milk. Over the summer, cases of campylobacteriosis continued, and on July 29, representatives from UDOH, UDAF, and UPHL conducted a collaborative investigation of the dairy specifically designed to detect the presence of Campylobacter, a fastidious organism very sensitive to pH. The bulk milk tank at the dairy was agitated, and a 1-liter sample

Utah Department of Agriculture and Food notified

14

No. of cases

12

Dairy inspected

10 8 6

Permit suspended

Permit reinstated

Permit permanently revoked

4 2 0 8 15 22 29 5 12 19 26 3 10 17 24 31 7 14 21 28 4 11 18 25 2 September May June July August

9 16 23 30 6 13 20 27 4 11 October November December

Illness onset week

Week of illness onset among patients (N = 99) with probable and confirmed Campylobacter jejuni infection associated with consumption of raw milk from a dairy—Utah, May–November 2014.



Case Study Exercise 12  Viable Plate Count

was obtained, adjusted to a pH of 7.5, and used to inoculate sheep blood agar, a rich growth medium. The number of total bacteria, coliform bacteria and somatic cells, all fell within acceptable limits, but Campylobacter jejuni was found growing in the milk, and the DNA profile of the bacterium matched that seen in the three initial patients. On August 4, the UDAF suspended Ropelato Dairy’s permit to sell raw milk. After retesting revealed acceptable bacterial, coliform, and somatic cell counts, along with a lack of Campylobacter, the UDAF reinstated the dairy’s raw milk permit on October 1. Unfortunately, 7 new cases of C. jejuni infection were seen

133

over the next 5 weeks, and the UDAF permanently revoked the dairy’s raw milk permit on December 1st. Although many people believe raw milk provides health benefits not available in pasteurized milk, there is no scientific evidence to support these claims. According to the Centers for Disease Control, unpasteurized milk is 150 times more likely to cause illness and results in 13 times more hospitalizations than pasteurized dairy products. When asked if raw milk is safe to drink, the Food and Drug Administration, the Centers for Disease Control, and the American Academy of Pediatrics all had the same answer. No.

CASE STUDY ANALYSIS 1. A common method of ensuring pasteurization of milk is to test the activity of an enzyme called phosphatase. An adequately pasteurized milk sample will have only a very small amount (